Glial Cell line-derived Neurotrophic Factor Effects on …etheses.bham.ac.uk/3268/1/Gale12PhD.pdfGlial cell line-derived neurotrophic factor (GDNF) is a growth factor promoting survival,

Glial Cell line-derived Neurotrophic Factor Effects on

Dental Pulp Cells and Osteoblast-like Cells

by

Zoe Gale

A thesis submitted to

The University of Birmingham

for the degree of

DOCTOR OF PHILOSOPHY

Department of Oral Biology

School of Dentistry

College of Medicine and

Dentistry

December 2011

ii

University of Birmingham Research Archive e-theses repository

This unpublished thesis/dissertation is copyright of the author and/or third parties. The intellectual property rights of the author or third parties in respect of this work are as defined by The Copyright Designs and Patents Act 1988 or as modified by any successor legislation. Any use made of information contained in this thesis/dissertation must be in accordance with that legislation and must be properly acknowledged. Further distribution or reproduction in any format is prohibited without the permission of the copyright holder.

iii

ABSTRACT

Glial cell line-derived neurotrophic factor (GDNF) is a growth factor promoting

survival, proliferation and differentiation of neural crest cells. Neural crest cells play an

important role within mesenchymal tissues during dental pulp and calvarial bone

development. GDNF also has a role within non-neuronal tissues and is expressed during

dental development. GDNF null mutations prevent the formation of the mineralised

hard tissues of the tooth. The hypothesis for this study was that GDNF affects

mesenchymal dental pulp cells (DPC), promoting the regenerative responses of

mineralised tissues. This study utilised cell culture models to investigate the direct

effects of GDNF on the proliferation and differentiation of dental pulp cells, bone

marrow mesenchymal stromal/stem cells (BMSC) and calvarial osteoblasts. This

research demonstrated that these culture models expressed GDNF and its receptors

GFRα1 and RET. GDNF was shown to directly stimulate DPC and osteoblast-like cell

proliferation and differentiation. Moreover, GDNF was cytoprotective when DPCs were

cultured under conditions reflecting aspects of inflammation, which may occur during

repair. These conditions included supplementation with the pro-inflammatory cytokine

TNFα and culture under serum-starved conditions. It is proposed that GDNF may play

an important regulatory role in dental pulp homeostasis and bone metabolism.

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For Erin, Shayla and Zena

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ACKNOWLEDGEMENTS

I would like to thank the University of Birmingham who funded this project, Professor AJ

Smith and the department of Oral Biology for allowing me to conduct this research within their

laboratories. I would also like to thank the staff of the Oral Biology laboratory; Gay, Sue and

Michelle for their helpful advice and support.

I have learnt so much under the supervision of Dr BAA Scheven and Dr PR Cooper, aside from

their expertise within mineralised tissue research, I would like to thank Dr Scheven for his

enthusiasm, precision, discipline and mentoring. I would like to thank Dr Cooper for his

dedication, guidance and strong work ethic.

I would also like to acknowledge staff and students within several departments of the College of

Medicine and Dentistry for their constructive critism, helpful advice and use of

reagents/equipment, particularly Dr Matthews, Dr Carter, Dr Wright, Dr White, Dr Shelton, Dr

Leadbeater, Dr Landini, Mr U Patel and Dr Wang.

I give a special thank you to Sue, Gay, Lisa, Erum and Vince for your genuine kindness and

sincerity.

Last not least, thank you to my husband Jon, who was also my IT/technical support service, my

children who are the love of my life and my closest family for their support and understanding.

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TABLE OF CONTENTS

LIST OF FIGURES

LIST OF TABLES

LIST OF ABBREVIATIONS

CHAPTER 1 INTRODUCTION

1. GDNF and Neurotrophic Growth Factors (NTGF).................................................1

1.1.1 GDNF receptors and signalling....................................................................2

1.1.2 GDNF, cell proliferation, survival and apoptosis........................................7

1.2. Tissue development and GDNF function...............................................................8

1.2.1 Tooth development ....................................................................................11

1.2.2 Primary, secondary and tertiary dentinogenesis.........................................15

1.2.3 Intramembranous and endochondral bone development ...........................17

1.2.4 Dentine and bone extracellular matrix (ECM)...........................................19

1.3 GDNF and repair of mineralised tissue.................................................................22

1.3.1 Mesenchymal stromal/stem cells (MSC) ……………………………......23

1.3.2 The role of nervous system in mineralised tissue repair............................27

1.3.3 The role of GDNF in mineralised tissue repair..........................................29

1.4. Aims of this study...................................................................................................34

CHAPTER 2 MATERIALS AND METHODS

2.1 Cell isolation...........................................................................................................37

2.1.1 Culture of bone marrow stromal/stem (BMSC) cells.............................................37

2.1.2 Dental pulp extraction ..........................................................................................37

2.1.3 Explant-derived dental pulp cells (DPC) isolation................................................38

2.1.4 Enzyme-derived DPC isolation..............................................................................38

2.1.5 Cell lines culture....................................................................................................40

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2.1.6 Cell culture............................................................................................................40

2.1.7 Cell counting .........................................................................................................41

2.1.8 Preparation of serum-free medium.........................................................................41

2.2 Preparation and storage of GDNF .........................................................................41

2.2.1 Preparation and storage of tumour necrosis factor alpha (TNFα)..........................43

2.2.2 Preparation and storage of nerve growth factor beta (NGF-β) ..............................43

2.2.3 WST-1 viable cell number assay ..........................................................................43

2.2.4 RET receptor inhibition (RPI-1)...........................................................................44

2.2.5 GFRα1 receptor inhibitor (PI-PLC).....................................................................44

2.2.6 BrdU (5-Bromo-2’-deoxy-uridine) assay...............................................................47

2.2.7 Live and dead assay ..............................................................................................47

2.2.8 Lactate Dehydrogenase (LDH) assay.....................................................................48

2.2.9 Caspase Glo3/7 assay ............................................................................................48

2.3 Streptococcus mutans (S.mutans) analysis.............................................................49

2.3.1 Mineralisation assay ..............................................................................................49

2.3.2 Alkaline phosphatase (AlP) assay .........................................................................52

2.4 Reverse transcriptase polymerase chain reaction (RT-PCR) analysis ..................53

2.4.1 Agarose gel electrophoresis and image analysis ...................................................55

2.5 Immunocytochemistry............................................................................................56

2.6 Statistical analysis .................................................................................................57

RESULTS

CHAPTER 3 CHARACTERISATION AND ANALYSIS OF

CULTURE MODELS

3. Characterisation of cell cultures ............................................................................60

3.1 DPC, BMSC and MC3T3-E1 cells standard curve analysis under varying

culture conditions...............................................................................................60

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3.1.1 Comparison of growth curves for explant-derived and enzymatically-

derived DPC under serum supplemented conditions...........................................62

3.1.2 DPC, BMSC and MC3T3-E1 cell viability under serum-free and serum-

supplemented conditions.....................................................................................64

3.2 Relative gene expression profiles in DPC, BMSC and MC3T3-E1

cells..................................................................................................................................69

3.2.1 GDNF, GFRα1 and RET receptor relative gene expression within a

glial cell line........................................................................................................69

3.2.2 Relative gene expression of GDNF, GFRα1 and RET in DPC.................70

3.2.3 Relative gene expression of GDNF, GFRα1 and RET in BMSC and

MC3T3-E1 cells..................................................................................................72

3.3 Immunocytochemical analysis of GDNF receptor expression ...........................73

CHAPTER 4 GDNF EFFECTS ON SURVIVAL AND PROLIFERATION

4 GDNF effects on viable cell numbers ......................................................................79

4.1 GDNF effects on cell proliferation and cell viability....................................83

4.1.1 Effect of GDNF on DPC and MC3T3-E1 cell proliferation......................84

4.2. Effect of GDNF on DPC, BMSC and MC3T3-E1 cell survival..................88

4.2.1 Effect of GDNF on cellular apoptosis and necrosis...................................95

4.3 The effect of attenuating GDNF signalling on DPC and MC3T3-E1 cell

viability...........................................................................................................................97

4.3.1 The effect inhibition of GFRα1 (GPI- linked receptor).............................97

4.3.2 The effect of inhibition of RET receptor DPC and MC3T3-E1 cell

viability...............................................................................................................99

CHAPTER 5 GDNF EFFECTS ON CELL

DIFFERENTIATION AND IN VITRO MINERALISATION

5 Short-term in vitro differentiation...........................................................................102

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5.1 Long-term in vitro mineralisation ..............................................................104

5.1.1 In vitro mineralisation under serum-free conditions................................107

CHAPTER 6 POTENTIAL ROLE OF GDNF DURING

REPAIR AND INFECTION

6 GDNF effects on DPC and/or MC3T3-E1 cell behaviour in response to;...........112

6.1 Streptococcus mutans (S. Mutans)...............................................................112

6.1.1 Tumour necrosis factor alpha (TNFα)......................................................115

6.1.2 Nerve growth factor-beta (NGF-β) ..........................................................119

6.2 Effect of GDNF on short term gene expression in DPC and MC3T3-E1 cells

.......................................................................................................................................122

CHAPTER 7 DISCUSSION

7 DPC culture model characterisation.......................................................................126

7.1 GDNF and GDNF receptor expression in DPC cultures ............................128

7.1.1 Confirmation of the mesenchymal nature of DPC culture model............129

7.1.2 Characterisation of bone cell cultures for GDNF and receptor

expression..........................................................................................................130

7.1.3 In vitro culture under serum-free conditions............................................131

7.2 Short term effect of GDNF on cell proliferation under serum-free

conditions.....................................................................................................................133

7.2.1 GDNF and cell survival ..........................................................................133

7.2.2 Osteogenic differentiation and in vitro mineralisation............................134

7.3 Short-term effect of GDNF on cell proliferation and differentiation under

serum-supplemented conditions.....................................................................136

7.4 In vitro culture under challenging conditions that reflect some aspects of

infection and inflammation.........................................................................................137

7.4.1 S.mutans exposure....................................................................................138

7.4.2 GDNF interaction with TNFα .................................................................139

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7.4.3 Cell specific effects of GDNF................................................................140

7.4.4 NGF stimulation of DPC........................................................................141

7.5 Summary ..............................................................................................................142

CHAPTER 8 CONCLUSION

8.1 Clinical and therapeutic applications......................................................................143

8.1.2 GDNF as a bioactive in tissue repair and regeneration ......................................143

8.1.3 Role of GDNF in tooth inflammation and analgesia..........................................145

CHAPTER 9 REFERENCES……...…………………………..…..….146

APPENDIX

List of publications, conference papers and abstracts..................................................199

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LIST OF FIGURES

Figure 1 Neurotrophins and glial family ligands receptor interactions..........................3

Figure 1.1 Intracellular signalling pathways activated by GFRα/RET.............................4

Figure 1.2 Schematic detailing the signalling involved during apoptosis ........................9

Figure 1.3 The gross anatomy of the developing embryo ..............................................12

Figure 1.4 Stages of rodent tooth development...............................................................14

Figure 1.5 Intramembranous ossification and endochondral bone formation ................20

Figure 1.6 Immunohistochemical localization of nerve fibres within bone....................28

Figure 1.7 Innervation of teeth........................................................................................31

Figure 1.7.1 Immunohistochemical localisation of nerve fibres and odontoblasts........32

Figure 2 Extracted dental pulp........................................................................................39

Figure 2.1 Enzymatically-derived and explant-derived DPC.........................................42

Figure 2.2 Trypan blue staining of DPC.........................................................................50

Figure 2.3 Linear standard curve for Alizarin red stain (ARS)......................................52

Figure 2.4 Negative controls for immunocytochemical staining....................................59

Figure 3 Explant-derived and enzymatically-dervied DPC standard curve (WST-1).....61

Figure 3.1 MC3T3-E1 cells and BMSC standard curve (WST-1)..................................62

Figure 3.1.1 Number of viable explant and enzymatically derived DPC at 48 hours.....63

Figure 3.1.2 Effect of serum-free culture on explant-derived DPC ..............................65

Figure 3.1.3 Effect of serum-free conditions on BMSC and MC3T3-E1 cells..............67

Figure 3.1.4 Effect of serum-free culture on the number of viable cells up to 10days..68

Figure 3.2 sq-RT-PCR analysis of C6 glioma cell line..................................................69

Figure 3.2.1 sq-RT-PCR analysis of explant-derived and enzymatically-derived

DPC.................................................................................................................................71

Figure 3.2.2 sq-RT-PCR analysis of BMSC...................................................................73

Figure 3.2.3 sq-RT-PCR analysis of MC3T3-E1 cells...................................................74

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Figure 3.3 Immunohistochemical staining C6 Glioma cells..........................................75

Figure 3.3.1 Immunohistochemical staining DPC.........................................................76

Figure 3.3.2BMSC immunohistochemical staining.......................................................77

Figure 3.3.3 MC3T3-E1 cell immunohistochemical staining........................................78

Figure 4 GDNF effect on C6 glioma cell number ........................................................80

Figure 4.1 GDNF effect on viable DPC number...........................................................81

Figure 4.1.1 GDNF effect on viable MC3T3-E1 cell number ......................................82

Figure 4.1.2 GDNF effect on viable BMSC number....................................................83

Figure 4.2 BrdU labelling of DPC under serum-free conditions ..................................85

Figure 4.2.1 BrdU labelling of DPC under serum-supplemented conditions................86

Figure 4.2.2 BrdU labelling of MC3T3-E1 cells...........................................................87

Figure 4.3 Live and dead analysis of DPC at 24 hours ................................................89

Figure 4.3.1 Live and dead analysis of DPC at 48 hours..............................................90

Figure 4.3.2 Live and dead analysis of MC3T3-E1 cells at 24 hours...........................91

Figure 4.3.3 Live and dead analysis of MC3T3-E1 cells at 48 hours..........................92

Figure 4.3.4 Live and dead analysis of BMSC at 24 hours ........................................93

Figure 4.3.5 Live and dead analysis of BMSC at 48 hours..........................................94

Figure 4.4 Effect of GDNF on LDH levels of DPC, MC3T3-E1 and BMSC.............95

Figure 4.5 Caspase 3/7 activity within DPC cultures...................................................96

Figure 4.6 PI-PLC effects on GDNF-stimulated DPC and MC3T3-E1 cells..............98

Figure 4.6.1 RPI-1 effect on GDNF-stimulated DPC viable and MC3T3-E1 cells....100

Figure 4.6.2 RPI-1 effect on GDNF-stimulated glioma cells......................................101

Figure 5 GDNF effect on DPC, MC3T3-E1 cells and BMSC AlP activities.............103

Figure 5.1 Effect of osteogenic culture conditions on in vitro mineralisation............106

Figure 5.2 Effect of GDNF on mineralised matrix formation for DPC cultures .......107

Figure 5.3 Effect of GDNF on in vitro DPC mineralisation.......................................110

Figure 5.4 Effect of GDNF on in vitro MC3T3-E1 cell mineralisation.....................111

Figure 6 GDNF effect on viable cell number with or without S. mutans .................114

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Figure 6.1 TNFα alone or GDNF and TNFα effect on viable DPC number ..............117

Figure 6.1.1 TNFα alone or TNFα and GDNF effect on viable MC3T3-E1 cell number

......................................................................................................................................118

Figure 6.2 NGF effect on viable DPC number............................................................120

Figure 6.2.1 NGF effect on AlP activity .....................................................................121

Figure 6.3 The effect of GDNF on DPC relative gene expression .............................124

Figure 6.3.1 The effect of GDNF on MC3T3-E1 relative gene expression ................125

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LIST OF TABLES

Table 1 Disease/dysfunction resulting from GDNF/GDNF receptor mutations..............6

Table 1.2 GDNF, RET, GFRα1, NGF and p75NTR within the developing tooth....35-36

Table 2.0 Summary of experimental assays and conditions used. ..........................45-46

Table 2.1 Details of primers and assay conditions for PCR analysis............................58

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ABBREVIATIONS

AA ascorbic acid or Vitamin C

ADM adrenomedullin

AKT acutely transforming retrovirus AKT8 in rodent T cell lymphoma

(also known as protein kinase B)

AlP Alkaline phosphatase

Apaf-1 apoptotic peptidase activating factor 1

ART artemin

ATM ataxia telangiectasia mutated

Bad Bcl-2-associated death promoter

Bak BCL2-antagonist/killer 1

Bax BCL2-associated X protein

Bcl B cell leukaemia

BDNF brain derived neurotrophic factor

β-gp β-glycerophosphate

BH3 Bcl-2 Homology (BH) domain 3

Bid BCL-2 Interacting Domain

BMP bone morphogenic proteins

BMSC bone marrow mesenchymal stromal/stem cells

BSP bone sialoprotein

casp caspase

CB Calbindin

Cbfα1 Core binding factor 1

CGRP calcitonin gene related peptide

Chk2 CHK2 checkpoint homolog (S. pombe)

Cn nerve cranial nerve five

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CNTF ciliary neurotrophic factor

Cyt cytochrome

D dentine

DEX dexamethasone

DMP-1 dentine matrix protein-1

DNA deoxyribonucleic acid

DPP dentine phosphoprotein

DPSC dental pulp stem cells

DSP dentin sialoprotein

DSPP dentin sialophosphoprotein

E embryonic day

ECM extracellular matrix

ERK extracellular signal-regulated kinase

FACS fluorescence activated cell sorting

FADD fas-associated protein with death domain

FGF fibroblast growth factor

G1 gap phase 1

G1/S gap phase1/synthesis phase

GAL galanin

GDNF glial cell-line derived neurotrophic factor

GFRα GDNF family receptors alpha

GPI glycosyl phosphostidylinositol

GRB2 growth factor receptor-bound protein 2

HGF hepatocyte growth factor

HIF1α hypoxic inducible factor 1alpha

HOX Homeobox

xvii

HtRA2 HtrA serine peptidase 2

IAPs inhibitors of apoptosis

IDE inner dental epithelial

IGFI insulin like growth factor I

IL-6 interleukin 6

JNK c-Jun, N-terminal kinase

LPS lipopolysaccharide

LTA lipoteichoic acid

MAP2-positive microtubule associated proteins 2

MAPK mitogen activated protein kinase

MDM2 Mdm2 p53 binding protein homolog (mouse)

Met c-met transforming gene proto-oncogene encoded

MGIF murine glial cell-derived neurotrophic factor inducible

transcription factor

MSC mesenchymal multipotent/stem cells

mTOR mammalian target of rapamycin

NCAM neural cell adhesion molecule

NCC neural crest cells

NF200 neurofilament 200kDa

NFĸB nuclear factor kappa B

NGF nerve growth factor

NGF-β nerve growth factor beta

NP neuropeptides

NPY neuropeptideY

NT-3 neurotrophin 3

NT-4/5 neurotrophins 4/5

xviii

NTN neurturin

OB odontoblast layer

Ob osteoblast

Oc osteoclast

OCN osteocalcin

ON osteonectin

OP osteopontin

p19ARF p19 alternative reading frame

p75NTR

low affinity nerve growth factor receptor

PAX paired box genes

PDGF platelet derived growth factor

PDK pyruvate dehydrogenase kinase

PI3-kinase phosphoinositide 3-kinase

PI-PLC phospholipase C

PKC protein kinase C

PLC-y phospholipase C-gamma

PP2A protein phosphatase 2A activator, regulatory

PSP persephin

PTEN phosphatase and tensin homolog

RAF v-raf-1 murine leukaemia viral oncogene homolog 1

RAS rat sarcoma protein

RET rearranged during transfection

RTK tyrosine kinase receptor

sCNS sympathetic central nervous system

SH3 Src homology 3

Shh sonic hedgehog

xix

SN subodontoblastic nerve plexus

SOS son of sevenless

SOX SRY (sex determining region Y)-box

SP substance P

Src sarcoma virus protein

STAT signal transducer and activator of transcription

S.mutans Streptococcus mutans

t trabaeculae

TCF-4 T-cell factor 4 (also known as TCF7L2)

TGF-β transforming growth factor beta

TIEG TGF-beta-inducible early-response gene

TNFα tumour necrosis factor alpha

TrKA tyrosine kinase receptor A

VEGF vascular endothelial growth factor

VIP vasoactive intestinal peptide

Wnt wingless-type MMTV integration site family

xx

1

CHAPTER 1 INTRODUCTION

Factors regulating the development of mineralised tissues may also be involved

in the subsequent repair and regeneration of the tissue. The primary focus of this

research was to investigate the possible roles of Glial Cell line-Derived Neurotrophic

Factor (GDNF) in the regulation of non-neuronal cells that form mineralised tissues, by

using odontoblast-like and osteoblast-like cell culture models.

1 GDNF and Neurotrophic Growth Factors (NTGF)

GDNF was first discovered in the medium of the B49 glial cell line and was

identified as a survival factor for dopamine secreting neurons (Lin et al. 1993). It is

secreted as a pro-form that is cleaved to give the mature growth factor, which is a

dimeric protein of approximately 33-45 kDa (Lin et al. 1994; Green 2002). GDNF

belongs to the cysteine knot growth factor super-family and the locations of most

cysteine residues within GDNF are homologous to those within the TGF-β2

(transforming growth factor-beta2) structure (Chen et al. 2000). The TGF-β family of

proteins along with other major classes of proteins such as BMPs (bone morphogenic

proteins) are considered to be members of the NTFG family. The GFLs (glial family

ligands) consists of GDNF, Neurterin, Artemin and Persephin and are regarded as a part

of the larger NTGF family which also includes the neurotrophins; Nerve Growth Factor

(NGF) and Brain Derived Neurotrophic Factor (BDNF), Neurotrophin 3 (NT-3) and

Neurotrophins 4/5, (NT-4/5) (Levy et al. 2005; Freund-Michel and Fossard 2008;

Sariola and Saarma 2003). GFLs and NTGFs are known survival factors for neurones

(Airaksinen and Saarma, 2002; Hirano et al. 2000; Bromberg and Wang 2009) and

includes groups such as the neurokines (e.g. IL-6) which can function during early

2

repair and development (Nibali et al. 2012; Heinrich et al. 2003). Family members

such as IL-6 are also known to mediate oral pathologies and diseases of bone and joints

(Nibali et al. 2012; Nishimoto and Kishimoto 2006; Kishimoto 2010).

Neurotrophins have been shown to stimulate the proliferation of numerous cell

types in culture, including neurones, liver and muscle cells (Kitomitsu 2000; Brodie and

Sampson 1987). However, in certain cases in vitro actions are not always replicated in

vivo, for instance, in vitro NGF stimulates the proliferation of muscle cells whereas in

vivo within damaged muscle, it appears not to have the same effect (Brodie and

Sampson 1987; Menetrey 2000). NGF affects cell survival and apoptosis via binding to

p75NTR and TrKA receptors (Schor 2005; Bredesen et al. 2005; Mizuno 2007).

1.1.1 GDNF receptors and signalling

GDNF dimers elicit intracellular signalling by binding to the GDNF family

receptors alpha (GFRα1) (glycosyl phosphostidylinositol (GPI) linked subunits) and

RET (Rearranged during Transfection) (Figure 1). RET dimerises, forming the core of

the complex with outer GFRα1 dimers (Airaksinen and Saarma 2002; Sariola and

Saarma 2003; Naughton et al. 2006). RET is a tyrosine kinase receptor (RTK) and can

also be classified as a dependence receptor that is able to initiate opposing signalling

cascades, i.e. either survival or apoptosis (Goldschneider and Mehlen 2010; Bredesen et

al. 2005; Mehlen 2005; Del Rio et al. 2007). RET binding is common to all GFL’s

however, these molecules bind more selectively to the GFRα receptor isoforms (Figure

1).

3

LIGANDS Receptors Interaction Receptors LIGANDS

GDNF

NGF

NTN

BDNF

ART

NT-3

PSP

NT-4/5

KEY

Most specific binding

Non-specific binding

Figure 1 Receptor interactions between neurotrophins (NT) and glial family

ligands (GFL). GDNF = glial cell-line derived factor; NTN = neurturin;

ART=Artemin; PSP=Persephin; NGF = nerve growth factor; BDNF= (brain derived

neurotrophic factor); NT-3= (Neurotrophin 3); NT-4/5= (Neurotrophins 4/5).

GDNF binding to RET and GFRα1 modulate cell survival, apoptosis,

proliferation and differentiation via activated intracellular signalling cascades (Figure

1.1). GDNF activation of RET dimers, results in autophosphorylation of the RET

receptors (Jing et al. 1996; Sariola and Saarma 2003; Manié et al. 2001) and leads to

activation of Ras, PLC-y (PKC), Ras/MAPK, JNK and PI3/AKT intracellular signalling

cascades (Airaksinen and Saarma 2002; Coulpier et al. 2002; Manié et al. 2001)

(Figure 1.1) (Airaksinen and Saarma 2002; Airaksinen et al. 1999).

Gfrα2

Gfrα1

Gfrα3

RET

P75NTR

Gfrα4

TrKA

TrKB

TrKC

4

Figure 1.1 Intracellular signalling pathways activated by Gfrα/RET

phosphorylation (adapted from Prazeres et al. 2011). Abbreviations; PI3-kinase,

phosphoinositide 3-kinase; PDK, pyruvate dehydrogenase kinase; HIF1α, hypoxic

inducible factor; VEGF, vascular endothelial growth factor; AKT, acutely transforming

retrovirus AKT8 in rodent T cell lymphoma or protein kinase B; mTOR, mammalian

target of rapamycin; STAT, signal transducer and activator of transcription; SH3, Src

homology 3; GRB2 =; SOS, son of sevenless; RAS, rat sarcoma protein; RAF, v-raf-1

murine leukemia viral oncogene homolog 1; ERK, extracellular signal-regulated kinase;

PLC-y, phospholipase C-gamma; JNK, c-Jun, N-terminal kinase; TCF-4, T-cell factor

4 (also known as TCF7L2).

Cell membrane

RPI-1

Extracellular receptor

portion

5

Interestingly the GFRα1 receptor may also be present as a soluble isoform and

can subsequently activate signalling in a paracrine manner (Airaksinen and Saarma

2002; Sariola and Saarma 2003). This type of signalling involving soluble GFRα1 with

RET may sustain GDNF effects (Manié et al. 2001). Furthermore, the GDNF-GFRα1-

RET complex may bind in a cell-to-cell manner, whereby one cell may express RET and

the other GFRα1 (Sprinzak et al. 2010). Poteryaev et al. (1999) found that GDNF and

Gfrα1 binding could occur independently of RET binding resulting in activation of

cAMP, MAPK and PLC-y intracellular signals. Later it was also determined that GDNF

signalling through GFRα1 without RET activates Src, a tyrosine kinase receptor (Brown

and Cooper 1996) and GDNF also activated Met, the Hepatocyte growth factor (HGF)

receptor, present on tumour cells (Sariola and Saarma 1999; Gao and Vande Woude

2005). Popsueva et al. (2003) indicated that direct Met and GDNF interactions did not

however occur within the kidney and therefore these findings suggest that RET-

independent actions of GDNF may be cell-type or tissue specific. NCAM (neural cell

adhesion molecule) is also a GPI linked receptor that may act as a GDNF co-receptor

mediating the mitogenic effects of GDNF within Sertoli cells (neural crest derived cells)

of the testes independently of RET (Yang and Han 2010). NCAM is expressed within

the developing dental pulp, however, unlike RET-GDNF-GFRα1 complexes, GDNF and

NCAM expression were not found to be overlapping during tooth development (Obara

et al. 2002) (see Table 1.2 and Figure 1.4). NCAM is expressed by differentiated

osteoblasts of human cranial bones (Marie et al. 2002) and during long bone

development, although NCAM expression appears more prominent during cartilage

formation compared with osteoblast formation (Hall and Miyake 1995). GDNF

signalling may also be dependent on cells being in contact with the extracellular matrix

6

(ECM), i.e. requiring cell surface-associated heparan sulphate glycosaminoglycans

(Lindfors 2006; Sariola and Saarma 2003). RET protein structure contains cadherin-like

and fibronectin binding motifs that are also observed within other adhesion molecules

indicating that RET may couple the cytoskeleton to the ECM (Kuma et al. 1993;

Anders et al. 2001; Gumbiner et al. 2005).

Interestingly GFRα receptors are much more widely expressed in the nervous

system compared with RET receptors (Trupp et al. 1997). Okragly and Haak-Frendscho

(1997) reported on levels of GDNF expression within the organs of postnatal mice,

finding that higher concentrations are found within the brain compared to the kidney and

other organs. The importance of the role of the GDNF receptor complex in neuronal and

non-neuronal tissue development is highlighted by observation of the phenotypes which

occur as a result of genetic mutations in GDNF and its receptors (Table 1).

Gene Phenotype Reference

RET Null mutation results in death at birth or

shortly after. Lack of neural crest derived

neuronal innervation of gut. Kidneys fail to

fully develop or may be absent.

Schuchardt et al.

(1994)

GFRα1 Null mutations die 24 hours after birth.

Reduction in neurons of the colon. No

kidneys develop.

Enomoto et al. (1998);

Garcès et al. (2000)

G GDNF Mutations cause megacolon (or

Hirschsprung's disease in humans). Null

mutations die at birth or shortly afterwards.

Lack of neural crest derived neuronal

innervation of gut. Dentine and enamel of

the teeth fail to develop.

Pichel et al. (1996)

De Vicente et al.

(2002)

Table 1. Disease/dysfunction resulting from gene mutations in GDNF and its

associated receptors in mice [adapted from Airaksinen et al. (1999) and Schor et al.

(2005)].

7

1.1.2 GDNF, cell proliferation, survival and apoptosis

GDNF acts as a pro-survival factor for a large number of cell types and is able to

exert mitogenic and differentiation effects (Weisenhofer et al. 2000; Ng et al. 2009;

Peterson et al. 2004; Toshifumi et al. 2005; Baldassarre et al. 2002 Meng et al. 2000;

Park et al. 2005; Wu et al. 2005 Insua et al. 2003; Hofmann et al. 2005; Mwizerwa et

al. 2011). Changes in cell cycle phases are linked with differentiation (Ohnuma et al.

2001; Buttitta and Edgar 2007) and studies have identified certain points within the cell

cycle that can be modulated by GDNF signalling, in particular G1 entry and G1/S phase

via p27kip1

and cyclin A regulation (He et al. 2008; Vitagliano et al. 2004; Baldassarre

et al. 2002).

Apoptosis plays an important role during tissue development, differentiation,

maintenance and repair as it allows excess or damaged cells to be safely removed

(Elmore 2007). For instance, during neuronal development excess cells are generated in

response to neurotrophins and those not receiving growth factor stimulation

subsequently apoptose (Yuan and Yankner 2000). Apoptosis and survival are controlled

by a balance between activators (pro-apoptotic proteins) and inhibitors (anti-apoptotic

proteins). Activation of the apoptotic process involves caspases-3, -6 and -7 and these

are termed effector or executioner caspases and are responsible for initiating both the

degradation of DNA and the end stages of apoptotic cascades (Nicholson 1999; Elmore

2007). Two major pathways of apoptosis are discerned and include the intrinsic or

mitochondrial pathway, whereby activators of apoptosis are released from the

mitochondria, and the extrinsic pathway, where apoptosis is induced following receptor

binding (Ashe and Berry 2003; Elmore 2007). Both processes ultimately result in DNA

8

and cytoskeletal degradation, nuclear fragmentation and the formation of apoptotic

bodies (Elmore 2007) (see Figure 1.2).

The extrinsic pathways leading to apoptosis can be influenced by growth factors

(see Figure 1.2, pg 9). GDNF and other growth factors can function as pro-survival

factors influencing the expression of anti-apoptotic factors such as Bcl family members

(Ghribi et al. 2001; Pugazhenthi et al. 2000). However, the intracellular domain of

RET has a caspase-3 cleavage site, which in the absence of GDNF signalling may be

cleaved to initiate apoptotic cascades (Bordeaux et al. 2000). GDNF signalling via RET

activates the AKT/P13 and Ras pathways that regulate cyclin D and stimulates cell cycle

progression, also GDNF stimulation suppresses BAD activity and up-regulates IAPs

(inhibitors of apoptosis) (see Figure 1.2) (Neff et al. 2002; Drosten et al. 2004;

Airaksinen and Saarma 2002). Similarly to the RET receptor, the TNFα receptor

(p75NTR

low affinity nerve growth factor receptor or CD271) can also supply both

positive and negative signals for survival determined by the activation of transcription

factor NFĸB that subsequently regulates apoptosis in response to TNFα (Wullaert et al.

2007; Liu et al. 1996).

1.2 Tissue development and GDNF function

Apoptosis occurs during cell differentiation within the developing calvarial

bones in order to prevent premature closure of the adjoined skull bones and this process

of cranial osteoblasts is proposed to be regulated by FGF-2 (beta-fibroblast growth

factor) and BMP-2 (Marie et al. 2002). Similarly, a wave of apoptosis has been

reported during tooth development that coincides with the differentiation of

odontoblasts (cells that form mineralized tissue of the tooth (see section 1.2.1 and 1.2.4)

9

(Mitsiadis et al. 2008) although the growth factors regulating this process have not

been reported. Organ development is often subject to reciprocal interactions, involving

epithelial tissue transmitting inductive signals (growth factors) and a mesenchymal

tissue being competent to respond to the signal by returning an inductive signal (Gurdon

1992; Thesleff et al. 1995; Zhang et al. 2005). In line with these reports, abnormal

GDNF expression and hence signalling between epithelial and mesenchymal tissues

within the developing lung has been linked with decreased levels of apoptosis and

subsequent

Figure 1.2 Schematic diagram detailing the signalling pathways involved

during apoptosis that may be regulated by growth factors such as GDNF (from

Johnstone et al. 2002). Component molecules in red inhibit apoptosis while those in

green promote apoptosis. Abbreviations used: FADD, Fas-Associated protein with

Death Domain; Apaf-1, apoptotic peptidase activating factor 1; Bid, BCL-2 Interacting

Domain; Smac, diablo homolog ; HtRA2, HtrA serine peptidase 2; Bax, BCL2-

associated X protein; Bak, BCL2-antagonist/killer 1; Bad, Bcl-2-associated death

promoter; Bcl-xL/Bcl-2 associated death promoter; PTEN, phosphatase and tensin

homolog; PP2A, protein phosphatase 2A activator, regulatory; BH3, Bcl-2 Homology

(BH) domain 3; ATM, ataxia telangiectasia mutatedChk2, CHK2 checkpoint homolog

(S. pombe); MDM2, Mdm2 p53 binding protein homolog (mouse); p19ARF

, p19

alternative reading frame, casp, caspase; cyt, cytochrome.

10

malformation of the organ structure (Fromont-Hankard et al. 2002). Epithelial-

mesenchymal signalling is a prominent process during tooth development and data

indicates that GDNF may play a role in such reciprocal interactions (Vainio and Lin

2002; Fromont-Hankard et al. 2002; Airaksinen 1999; Nosrat et al. 2002; Lukko et al.

1997; Nosrat et al. 1998; Fried et al. 2000).

Organ and tissue development reportedly occurs due to the actions of

morphogens (growth factors) forming concentration gradients that specify cell fates

within the embryo (Gierer and Meinhardt 1972; Green 2002). Notably GDNF as well as

FGF, TGFs and activin (a NTGF) have been identified as morphogens involved in tissue

development (Armelin 1973; Todaro et al. 1981; Vale et al. 1986; Slack et al. 1987;

Smith et al. 1990; Green and Smith 1990; Green 2002). These growth factors

subsequently influence a range of transcription factors such as HOX (Homeobox) genes,

PAX (Paired box genes) and SOX [SRY (sex determining region Y)-box genes] gene

families, specifying cell fates. Interestingly GDNF regulates SOX10 transcription

within neural crest cells (NCC), and SOX10 with PAX3 also regulate the transcription

of RET (Lang et al. 2000, 2003; Linnarsson et al. 2001; Wu et al. 2008).

During early embryogenesis NCC are generated from ectodermal tissue of the

mid- and hind-brain neural folds (Lumsden 1986, 1988; Ruch et al. 1995). These

pluripotent NCC migrate from the ectoderm throughout the embryo to specific sites

where they proliferate and differentiate to form structures within organs and tissues, e.g.

odontoblasts of the dentine-pulp complex of the tooth (Gilbert 2000; Nanci 2003;

Gronthos and Brahim 2002). Embryonic NCC develop into the adult cranial and sensory

neurones (including nocioceptors) (Nanci 2003), sympathetic neurones and

parasympathetic neurones (White and Anderson 1999), the periodontal ligament,

11

alveolar bone, the adrenal medulla, neurones of the gut (Kruger et al. 2002) cranio-

facial bones, the skull (Nanci 2003), Schwann cells (Stemple 1992; Kaltschmidt et al.

2012), glia (Henion et al. 2000), spinal ganglia (Lefcort and George 2007), carotid body

and aorta (mechanosensors) of the heart (Izal-Azcárate 2008, Gilbert 2000) (Figure 1.3).

This ability of ectodermally derived cells to migrate, inhabit and function as

mesenchymal cell types is unique to these pluripotent NCC.

GDNF affects developing NCC derived tissues, influencing cellular

proliferation, migration and differentiation throughout the embryo (Britsch et al. 2001;

Young et al. 2001; Natarajan et al. 2002; Mwizerwa et al. 2011). GDNF is also

expressed in postnatal neural crest derived tissues including the dermis (Motohashi

2006; Oshima et al. 2010), carotid body of the heart (Izal-Azcárate 2008; Shakhova and

Sommer 2010; Miwa et al. 2010; Leitner et al. 2005), the Sertoli cells of the testes

(Sariola and Immonen 2008), the inner ear (Stöver et al. 2001; Nam et al. 2000;

Fransson et al. 2010; Minoux and Rijli 2010) and retinal cells of the eye (Hauck et al.

2006; Grocott et al. 2011). It is now proposed here that GDNF may influence NCC

derived dental pulp and bone cell differentiation.

1.2.1 Tooth development

The pathway whereby NCC form odontoblasts begins with cranial NCC

migration from the cranial region of the embryo into the mesenchymal tissue of the

branchial arches. The first branchial arch (Figure 1.3, pg 12) develops into the jaw and

after cranial NCC migration into the branchial arch, the adjacent epithelium thickens

and invaginates into the NCC containing condensing mesenchyme (consisting of

aggregates of mesenchymal cells) to form a dental placode (Figure 1.4, pg 14). This

12

developing tooth subsequently proceeds through a series of developmental stages

termed bud, cap, bell and crown (see Figure 1.4, pg14).

During the bell stage the condensing mesenchyme develops into the dental

papillae and the epithelium forms the dental or enamel organ (Figure 1.4, pg 14)

simultaneously re-iterative signals (of growth factors) occur throughout the different

stages of tooth development, beginning with initial stimulation from the oral epithelium

to the cranial NCC derived mesenchyme (Thesleff and Tummer 2009). The growth

factor signalling involved reportedly includes BMPs, FGF, Wnt (wingless-type MMTV

integration site family) and Shh (sonic hedgehog) (Thesleff and Tummers 2009).

A. B.

Figure 1.3 (A) The gross anatomy of the developing human embryo showing the

position of the four branchial arches at the head end of the embryo. Red arrows

indicate branchial arch positions. (B) Section through the branchial (pharyngeal

arch). The red line indicates the saggital plane from where the cross section is derived.

The branchial arches are shown in sagittal plane as regions of the ectoderm, mesoderm

and endoderm. There would appear a mirror image of this structure on the right hand

side of the existing image if the whole cross section was shown, from

http://rad.usuhs.edu/medpix.

Branchial arches 1 2 3 4

A Branchial

arch

Point of

contact

between

ecto-and

endoderm

layers

(Homma ) al

Ectoderm layer

(Outermost epithelia) Mesoderm (layer) of

Cranial NCC

Endoderm layer

13

Migration and recombination studies have demonstrated that odontoblasts

develop from the NCC derived mesenchyme of the dental papilla and only form

following interaction between cranial NCC and branchial arch epithelium (Lumsden

1988; Thesleff et al. 1995; Zhang et al. 2005). The importance of correct control of

cranial NCC migration is highlighted in Di George syndrome where cranial NCC

migration is disrupted, subsequently branchial arch structures do not develop normally

and tooth development is therefore abnormal (da Silva Dalben et al. 2008).

Interestingly mutations in VEGF (vascular endothelial growth factor) cause the

phenotypic changes seen in Di George patients (Campbell 2003). Notably Tufro et al.

(2007) reported that VEGF activates RET and induces GDNF up-regulation, indicating

an interesting interrelationship between these molecules that may affect craniofacial

development. Furthermore, inhibition of GDNF-RET signalling may repress tooth

formation (Klein et al. 2006; Taketomi et al. 2005). Studies have previously shown

that GDNF mediates the differentiation of NCC during the development of several other

organ systems (Vainio and Lin 2002; Sariola and Saarma 2003).

Odontoblasts differentiate from the cells of the dental papillae and begin

producing pre-dentine (unmineralised collagenous ECM of dentine) during the late bell

stage. Differentiating dental papilla/pre-odontoblasts also produce proteins such as

fibronectin and collagen type III that are associated with the forming basement

membrane positioned between the dental papilla and the inner epithelial (IDE) (Smith

and Lesot 2001; Tzaifas and Kondados 2010; Ruch et al. 1982). Thesleff et al. (1978)

reported that both mesenchymal dental papilla cells and IDE cells are needed to produce

the basement membrane and this interaction initiated odontoblast differentiation.

14

Figure 1.4 Stages of rodent tooth development (adapted from Thesleff and Tummers

2009; Lumsden 1988). E=embryonic murine day; ODE= Outer dental epithelium;

IDE= inner dental epithelium; DP =dental papilla; ERM= epithelial cell rests of

Malassez; HRM= Hertwig's epithelial root sheath.

E10.5 E13.5

E17.5

E19

IDE

DP

E15.5

ODE

15

The basement membrane proteins fibronectin and decorin have previously been

shown to support and signal odontoblast polarisation and differentiation (Ruch et al.

1995; Ruch 1998) however more importantly growth factor signalling is considered

crucial to this process. In particular, IGF, FGFs (a and b), TGF-βs (-1, -2, -3) and BMP

(-2, -4, -6) have been identified as putative factors involved. Other growth factors that

may regulate odontoblastic differentiation include NGF, PDGF (platelet derived growth

factor) (Yokose et al. 2004), HGF (Ye et al. 2006), Gdf11 (growth/differentiation

factor 11) (Nakashima et al. 2002) and ADM (adrenomedullin) (Musson et al. 2010).

1.2.2 Primary, secondary and tertiary dentinogenesis

Dentine is located beneath the tooth enamel (Frank 1999) (Figure 1.7 pg 31) and

is initially synthesized during development in a process termed primary dentinogenesis.

Following tooth eruption dentine is continually formed at the periphery of the dental

pulp throughout the life of the tooth (Linde 1989) and this process is termed secondary

dentinogenesis (Linde 1989; Sloan and Smith 2007). If the dentine becomes damaged

after eruption further dentine may be produced as a replacement in a process referred to

as tertiary dentinogenesis (Smith and Lesot 2001; Smith 2001; Tziafas et al. 2000).

Odontoblasts have processes which are the first point of contact between the

peripheral dentine and dental pulp. Cellular processes from an odontoblasts extend

within the dentine tubules along with neighbouring nerve fibres potentially up to the

dentine border with the enamel (Frank 1999; Karjalainen and Le Bell 1987; Moss et al.

2005). Consequently, physiological changes within the dentine/enamel are detected by

the odontoblast and nerve processes (Chaussain-Miller et al. 2006; Smith et al. 2001;

Smith 2001; Karjalainens and Le Bell 1987) and may stimulate tertiary dentinogenesis.

16

Physiological changes in the biofilm affect the dentine and enamel. Teeth develop a

naturally occurring coating of bacteria termed a biofilm which can become excessively

populated by acid producing bacteria, such as Streptococcus mutans, that subsequently

cause tooth mineral dissolution releasing the components of the dentine/enamel

structure (Marsh 2006; Verstraeten et al. 2008). As this dental caries progresses the

odontoblast may also come into direct contact with the bacteria or its components.

Odontoblasts have been shown to express toll-like receptors (TLR) that bind bacterial

surface proteins and these interactions may initiate an immune and inflammatory

response within the dental pulp, which may also contribute to tertiary dentinogenesis

depending on the severity of the infection (Cooper et al. 2010; Berkovitz 2002; Jones

and Boyde 1977; Hirao et al. 2009; Botero et al. 2006; Horst et al. 2009; Durand et al.

2006).

Dentine production by existing odontoblasts is termed reactionary

dentinogenesis and is a form of tertiary dentinogenesis. Severe disease/trauma may

result in odontoblast apoptosis or necrosis (Mitsiadis et al. 2008), however

neighbouring post-mitotic terminally differentiated odontoblasts cannot divide to

produce new daughter cells to repopulate and repair the injury site (Berkovitz 2002;

Moss et al. 2005). Generation of new odontoblast-like cells, differentiating from

mesenchymal dental pulp stem/progenitor cells may occur under appropriate conditions

(see section 1.3.1) and dentine produced during this form of tertiary dentinogenesis is

termed reparative dentinogenesis (Smith and Lesot 2001; Sloan and Smith 2007; Shiomi

2000).

17

1.2.3 Intramembranous and endochondral bone development

Long/limbs bones are formed by endochondral ossification whereby cartilage is

formed initially and subsequently replaced by bone produced by osteoblasts. Osteoblasts

are differentiated from the mesenchyme surrounding the cartilaginous tissue. However,

the flat bone of the cranium develops primarily by intramembranous ossification,

whereby osteoblasts directly differentiate from cranial NCC-derived condensed

mesenchyme (Figure 1.5, pg 20). Unlike tooth and craniofacial bones, the limb bud

mesenchyme is not derived from cranial NCC (LeLièvre and Douarin 1975; Noden,

1988; Chai et al. 2000; Jiang et al. 2002; Couly et al. 1992, 1993; Lumsden et al.

1986). In particular, the frontal area of the cranium forms from cranial NCC and the

posterior region from mesoderm (Opperman et al. 1993, 1996; 2000; Couly et al. 1992,

1993; Santagati and Rijli 2003). Thus, due to the differences in cellular origins, the

fundamental developmental processes as well as behaviour of osteoblasts within cranial

and long bones may be different (Ornitz and Marie 2002; Depew et al. 1999) (Mao et

al. 2006). Indeed Moore et al. (2002) noted that fibroblast growth factor -2 (FGF-2)

implanted into the developing skull had no detectable effect yet when it was implanted

into the developing limb it initiated growth of this structure. Furthermore, null mutation

of the BMP receptor type 1B gene resulted in defects in limb bones but not cranial bone

formation (Lian et al. 2006) underscoring physiological differences between

endochondral and intramembranous ossification. Van der Bos et al. (2008) also reported

differences in levels of collagen, osteopontin and osteonectin within the ECM

composition of each bone type.

18

Little is known about the role of GDNF in bone development and osteoblast

differentiation. It is postulated here that GDNF may be involved in cranial bone

formation through a direct action on NCC. During intramembraneous ossification

mesenchymal cells condense, then differentiate into osteoprogenitors and osteoblasts

directly (Hall and Miyake 1992) (Figure 1.5). Osteoprogenitors and pre-osteoblasts

secret extracellular matrix (ECM) components (e.g., collagen III, hyaloronan, collagen

type I) that may be stimulated by growth factors such as TGF-β and these ECM

molecules may play a role in inducing further osteoblast differentiation (Lian and Stein

2003, 2004, 2006). Furthermore, autocrine secretion of BMP-2, BMP-4, and BMP-7 by

differentiated calvarial osteoblasts may have further effects stimulating cyto-

differentiation, increasing the transcription of bone constituents including osteocalcin

(OCN) and bone sialoprotein (BSP) transcripts (Xiao et al. 2002; van der horst et al.

2002).

The role of GDNF in long bone formation is also unclear. During long bone

formation, the proliferation and differentiation of osteoblasts is due to a series of

interactions between chondrocytes and a pre-osteoblast mesenchymal region (called the

perichondrium) (Vortkamp et al. 1998) secreting growth factors such as BMP-2, -4, and

-7 (Yamaguchi et al. 2000; Vortkamp et al. 1998), FGF-7, -8, -17 and -18 (Ornitz and

Marie 2002; Ducy et al. 1997; Yamaguchi et al. 2000). Later osteoblasts differentiate

from the mesenchyme (the perichondrium) surrounding the chondrocytes. GDNF

mRNA has been detected in mesenchymal tissues surrounding the forming cartilage in

femur limb buds at embryonic day 14.5 (Hellmich et al. 1996). Furthermore GDNF was

secreted by chondrocytes stimulated with TNFα (Kashani et al. 2010). Interestingly,

HOX11 null mice have no detectable expression of GDNF or the GDNF transcription

19

factor six2 (Wellik et al. 2002; Brodbeck et al. 2004). HOX11 functions together with

HOX10 (Myers 2008) and HOX9 to regulate digit formation in forelimbs (Xu and

Wellik 2011; Goff and Tabin 1997) and HOX11 mutant mice have spine and limb joint

abnormalities (Small and Potter 1993; Davis et al. 1995). This data suggests that

HOX11 is involved in transcription of GDNF during endochondral ossification.

1.2.4 Dentine and bone extracellular matrix

The mineralised matrix of dentine and bone consists of hydroxyapatite crystals

[Ca5(OH)(PO4)3] that are embedded in a collagenous ECM. The ECM affects the

position, orientation, size and growth of the crystal hydroxyapatite in these hard tissues

(Boskey 1991). Bone and dentine also contain non-collagenous proteins, including

phosphoproteins, dentine matrix protein-1 (DMP-1), dentine sialoprotein (DSP), dentine

phosphoprotein (DPP), bone sialoprotein (BSP), osteonectin (ON), osteocalcin (OCN),

osteopontin (OPN) and proteins from serum (such as growth factors and albumin)

(Smith et al. 2011; Hao et al. 2009; Huang et al. 2008). The acidic non-collagenous

phosphoproteins of dentine and bone are suggested to be the sites of nucleation for

hydroxyapatite formation on the ECM due to their capacity to bind to collagen and

calcium ions (Fujisawa and Kuboki 1991; Houlle et al. 1997; Saito et al. 1997; Boskey

1991; He et al. 2003).

The enzyme alkaline phosphatase (ALP) has been suggested to play an important

role in the ECM mineralisation process (Beertsen and van den Bos 1989). It was

hypothesized that ALP is involved in the generation of inorganic phosphate ions

necessary for matrix mineralisation whilst inhibiting factors that reduce inorganic

portions of the ECM (Beertsen and van der Bos 1992; Damek-Poprawa et al. 2006).

20

Figure 1.5 Diagrammatic comparison of human endochondral bone formation (A)

and intramembraneous ossification (B), from Orntiz and Marie (2002). Note the limb

bud and mesenchymal condensations initiation of bone development differs,

osteoprogenitors (pink) are formed directly from mesenchyme (clear) during

intramembranous ossification, these differentiate to osteoblasts (yellow) that later reside

on the inner lining surface of the bone facing the bone marrow cavity or in the

outermost periosteum lining of the bone surface (B), compared to (A) endochondral

ossification where chondrocytes (light blue) form directly from mesenchyme (clear).

Notably, bone ALP (tissue non-specific ALP or TNSALP) is used as a biochemical and

clinical marker for osteoblast activity and total serum ALP is used for biochemical

diagnosis of bone diseases (Singer and Eyre 2008; Karjalainens and Le Bell 1987).

Recently, mutations in human TNSALP genes and TNSALP null rats have been shown

to produce hypophosphatasia a bone disease characterized predominantly by decreased

skeletal mineralization (Whyte 2010), indicating ALP is required for the mineralisation

21

of bone. In vitro ALP has also been used as an early, albeit nonspecific, marker for

osteogenic cell differentiation and has been suggested to reflect in vitro mineralization.

During osteoblast differentiation, confluent osteoblast cultures are able to form

multilayers which coincide with a decrease in cell proliferation whilst the expression

levels of ALP increase; notably as the ECM matures during mineralised nodule

formation ALP levels subsequently decrease (Bellows et al. 1991; Porter et al. 2003;

Golub and Boesze-Battaglia 2007). In vitro mineralisation can be stimulated by the

presence of osteogenic supplements, β-glycerophosphate (β-gp), dexamethasone (DEX)

and ascorbic acid (AA or Vitamin C). β-gp is necessary for the formation of mineralised

collagenous matrix (Fratzl-Zelman N et al. 1998), supplying ―active‖ phosphate for the

mineralisation of the collagenous ECM and is also a substrate for ALP (Fratzl-Zelman et

al. 1998). Collagen synthesis is dependent on the presence of AA (Murad et al. 1983;

Fratzl-Zelman et al. 1998) and DEX is a pleiotropic synthetic steroid that is used as an

inducer of osteogenic differentiation in vitro and also increases ALP expression (Boskey

and Roy et al. 2008; Alliot-Licht et al. 2005; Cheng et al. 1994, 1996).

The ―master‖ gene regulators of transcription during osteoblast differentiation

are regarded as Osterix and Cbfα1/RUNX2 (Ducy et al. 1997). The processes involved

in the regulation of odontoblast differentiation are however less well understood, but,

similar transcriptional controls as to those in osteoblasts may be involved in regulating

this process (Narayanan et al. 2001; Hao et al. 2005; Qin et al. 2001, 2004, 2007).

Interestingly the DSPP (dentin sialophosphoprotein) promoter has putative binding sites

for Cbfα1 (Chen et al. 2005) and this gene encodes two proteins DSP and DPP, both of

which have been associated with odontoblast differentiation and dentinogenesis (Shi

2001).

22

1.3 GDNF and repair of mineralised tissue

Serum proteins such as albumin, PDGF, TGF-β, bFGF, IGFI and II and VEGF

have been identified within the dentine ECM, sequestered there during dentinogenesis

(Roberts-Clark and Smith 2000; Linde and Goldberg 1993; Finkleman et al. 1990;

Kinoshita, 1979). Similarly, during bone development growth factors are also

sequestered within the developing mineralised matrix and are involved in osteoclast-

osteoblast signalling and the regulation of bone remodelling (Hauschka et al. 1986).

These signalling molecules may also be present in the soft extracellular

stromal/mesenchymal matrix postnatally. Bone remodelling repairs small areas of

damaged bone as a part of the normal homeostatic postnatal mechanism, controlling

bone size, strength and density. However, research investigating postnatal repair

processes of mineralised tissues has suggested larger repairs utilise mechanisms that

mimic the developmental processes forming mineralized tissues (Robling et al. 2006;

Ferguson et al. 1998, 1999; Vortkamp et al. 1998 Tsiridis and Giannoudis 2006;

Silkstone et al. 2008; Smith and Lesot 2001; Simon et al. 2009, 2010). Several growth

factors such as BMP-2, PDGF, VEGF and TGF-β involved in embryonic tooth

development and bone development are already known to influence postnatal dental

pulp physiology (Kinoshita 1979; Kato 1988; Lynch et al. 1999; Roberts-Clarke and

smith 2000; Tziafas et al. 2000; Matsushita 2000; Haddad et al. 2003; Kurihara 2003,

2007; Iohara et al. 2004; Sloan & Smith 2007; Scheven et al. 2009; Hauschka et al.

1986 ).

During injury/infection degradation of the mineralized ECM may liberate growth

factors and other proteins previously sequestered there (Chaussain-Miller et al. 2006),

where they may be bound to carrier molecules which maintain their stability, such as

23

heparin sulphate, decorin and biglycan (Berkovitz 2002; Karjalainens and Le Bell 1987;

Smith et al. 2011; Buckwalter and Mankin 1988). The release from the ECM following

disease may be key to signalling repair (Smith and Lesot 2001; Hing 2004), by evoking

migration, proliferation, survival and differentiation of osteo/odontoblasts, progenitors

and stem cells that can regenerate the mineralised tissue (Smith et al. 1990; 2001;

Benezra et al. 1993; Farges et al. 2003; Grassinger et al. 2009). However, the

existence of GDNF, NGF or other NTGFs among sequestered growth factors within

dentine is as yet unreported.

1.3.1 Mesenchymal stem/stromal cells (MSC)

Mesenchymal stem cells (MSC) have been identified within various

mesenchymal tissues including bone marrow, adipose and skin (Zuk et al. 2002),

kidney (Huang et al. 2009), umbilical cord (Lu et al. 2006) and muscle (Jankowski et

al. 2002). Neural crest stem cells within the mesenchyme such as the dental pulp may

also be considered as mesenchymal stem cells often referred to as ―ecto-mesenchymal‖.

The term stem cell is indicative of the self renewing or colony forming ability of cells

along with pluri-/multi-potentiality. MSC may be recruited to sites of injury, migrating

and differentiating to form tissue specific cells in situ (Koç et al. 2000; Crisan et al.

2011; Wise and Ricardo 2012; Theise et al. 2000; Peterson et al. 1999; Barry and

Murphy 2004). Distinct MSC populations can be obtained by FACS, however

significant work still remains to identify definitive panels of markers for MSC isolation

for all tissues (Gronthos et al. 2011; Battula et al. 2009; Kagami et al. 2011; Bühring

et al. 2007, 2009). The role of mesenchymal stem cells in tissue repair and regeneration

has generated a significant amount of interest over the last decade, in particular dental

pulp stem cells (DPSC) have now been considered as a potential and promising source

http://www.ncbi.nlm.nih.gov/pubmed?term=%22Buckwalter%20JA%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/pubmed?term=%22Buckwalter%20JA%22%5BAuthor%5D

24

for repair and regeneration of the tooth as well as other tissues (Sasaki et al. 2008; Nesti

et al. 2011; Gandia et al. 2008; Arthur et al. 2009; Kerkis et al. 2006, 2008; Zhang et

al. 2008). Although the precise identity and tissue localisation of DPSC are not fully

known, DPSC share many similarities with bone marrow MSC [(BMSC) bone marrow

mesenchymal stem/stromal cells] (Yan et al. 2011; Alge et al. 2010; Huang et al.

2009). DPSC and BMSC are multipotent, able to differentiate along osteogenic,

odontoblastic, adipogenic, neurogenic and chondrogenic lineages (Arthur et al. 2008;

Egusa et al. 2005; Sanchez-Ramos et al. 2000; Tziafas and Kondonas 2010). Similarly

to dentine regeneration, bone can regenerate during repair by osteoblast differentiation

from MSC (Hing 2004; Yu et al. 2007; Batouli et al. 2003). Thus, DPSC and BMSC

cultures may be used as model cultures pertaining to study of stem cells and the

processes involved in cell differentiation and tissue repair (Aubin 1998, 1999; Frank et

al. 2002; Zou et al. 2008; Wang et al. 2010b & c; Gronthos et al. 2002; Shi et al.

2005; Miura et al. 2003, 2004; Couble et al. 2000; Woodbury et al. 2002; Egusa et al.

2005).

The ability of MSC to differentiate along different lineages is influenced by

growth factors present within the injured/repairing tissue (Hermann et al. 2004; Barry

and Murphy 2004; Banfi et al. 2000) and MSC themselves also secrete a range of

growth factors that further influence the tissue repair and also act in an auto-/paracrine

manner (Majumdar et al. 1998; Barry and Murphy 2004). Interestingly, DPSC and

BMSC cultures produce a range of NTGFs including GDNF which prompted an

emerging interest in the use of these cells for neurological disorders and nerve repair

(Nosrat et al. 2001; Apel et al. 2009; Chop and Li 2002; Sasaki et al. 2008; Nesti et al.

2011). In addition, GDNF has been reported to induce stem cell self-renewal and to be

25

cytoprotective for MSC of the kidney stimulating proliferation and migration (Shi et al.

2008; Meng et al. 2000; Gonzalez et al. 2009). Therefore elucidation of the basic

functions of GDNF that control mesenchymal cell differentiation has potential for

broader beneficial applications.

Furthermore, GDNF effects may be co-ordinated with other growth factors such

as Wnts, TGF-βs, BMPs or mediators of inflammation, such as TNFα, neuropeptides

and neurotrophins, that are up-regulated during mineralised tissue repair (Golub and

Boesze-Battaglia 2007; Liu et al. 2008; Rawadi et al. 2003). Blocking Wnt signalling

abrogates ALP activity and results in abnormal tooth development (Qiang et al. 2008

Golub and Boesze-Battaglia 2007; Liu et al. 2008). Interestingly, Wnt signals down-

regulate GDNF, inhibiting stem cell self renewal during spermatogenesis (Tanwar et al.

2010) while Wnt11 deficiency decreases GDNF expression within the kidney (Chi et al.

2004). TGF-3 has also been reported to be required for recruiting the GDNF receptor

Gfr1 to the plasma membrane (Peterziel et al. 2007; Sariola and Saarma 2003) and

TGF-βs are required for the trophic actions of GDNF in vitro and in vivo (Krieglstein et

al. 2002; Schober et al. 1999). TGF-s are expressed in odontoblasts (Linde 1989,

Tziafas et al. 2000) and dental pulp cells, and isoforms I and III increased odontoblast

secretory activity, differentiation and repair in rat organ and cell culture models (Sloan

et al. 2002; Sloan and Smith 1999; Tziafas et al. 1998; Nie et al. 2006). Moreover

TGF-1 application within pulp capping experiments in vivo (Tziafas et al. 1998, 2001;

2000; Smith 2001) along with the inhibition of reparative dentinogenesis caused by

TGF-1 anti-sera application in ferrets (Smith et al. 1994) indicates TGF-1’s potential

important role in promoting dental hard tissue formation. Conceivably, differentiation

26

and mineralization events which occur during bone and dental pulp repair may involve

interactions between TGF-β and GDNF.

TNFα is a cytokine involved in immune and inflammatory responses at sites of

disease and injury (Szlosarek and Balkwill 2003; Konisti et al. 2012). TNFα inhibits

osteoblast terminal differentiation and has been found to be abundantly expressed in

many diseases of bone, including rheumatoid arthritis (Boyce et al. 2005). Dental pulp

cell TNFα expression is up-regulated by bacterial LPS (lipopolysaccharide) and LTA

(Lipoteichoic acid) and within infected pulpal tissue (McLachlan et al 2004; Bletsa et al.

2006; Keller et al. 2010). The role of TNFα in vivo during dental pulp inflammation

may however have opposing effects on DPC behaviour due to its potential ability to

promote repair processes at relatively low concentrations while inhibiting repair

processes at high concentrations (Cooper et al. 2010, 2011; Okabe and Matsushima

2006; Paula-silvia et al. 2009; Pezelj-Ribaric et al. 2002). Notably the GDNF promoter

has a binding site for NFĸB which is a key transcription factor activated in response to

TNFα signalling (Grimm et al. 1998) and GDNF expression in Sertoli cells is also

known to be up-regulated by TNFα stimulation (Simon et al. 2007; Hofmann 2008).

GDNF expression in the brain is also up-regulated during inflammation and repair; a

finding similar to that reported for GDNF within other non-neuronal tissues (Eberling et

al. 2009; Cheng et al. 2008; Katoh-Sembah et al. 2007; Nam et al. 2000; Tsui et al.

2006) indicating that GDNF specifically functions during soft tissue injury, infection or

repair. GDNF may therefore also influence hard mineralised tissue repair, directly or

indirectly via interactions with prominent growth factors and cytokines that regulate

these processes.

27

1.3.2 The role of the nervous system in mineralised tissue repair

Several functions have been ascribed to the up-regulation of NTGF and

neuropeptides (NP) released from the nervous system during injury/infection. Such

functions include the further stimulation of the nervous and immune systems to remove

infection, produce anti-inflammatory factors, modulate pain and perform neuronal repair

(Toben et al. 2011; Xian and Zhou 2009; Paschalia et al. 2011; Lundy and Linden

2004; Wang et al. 2003; Pezet and McMahon 2006; Lambrecht 2001). Highly

innervated bone (See Figure 1.6, pg 28 for the localisation of nerves) contains numerous

sympathetic nervous system fibres that have previously been suggested to regulate

haemopoietic cell proliferation and mobilisation from within the bone (Afan et al.

1997; Elmquist and Strewler 2005; Chen et al. 2002; Marenzana and Chenu 2008; Serre

et al. 1999; Delgado et al. 2004). However, more recently the innervation has been

suggested to be involved in bone development (Elmquist and Strewler 2005). This

newer more direct role of the sympathetic bone innervation and neuropeptides (NP) on

bone formation and putatively on repair was initially demonstrated by adrenergic

receptor activation, (Elefteriou 2005, Elefteriou et al. 2005; Elmquist and Strewler

2005) is currently under investigation (Lundy and Linden 2004; Sato et al. 2007; Goto

et al. 2007; Natsume et al. 2010). However NP are also secreted from non-neuronal

tissues during tissue damage and infection and these molecules can stimulate growth of

cells within mineralised tissues; for instance VIP (Vasoactive intestinal peptide), and

CGRP (calcitonin gene related peptide) are secreted by DPC, furthermore CGRP and SP

(substance P) were secreted by DPC in vitro after incubation with proteases from

P.gingivalis (Tancharoen et al. 2005). Both CGRP and SP also increased dental pulp

28

Figure 1.6 Immunohistochemical localization of neurones and nerve markers in long bones of growing rats aged 15-25 days. (a) Metaphysis of

neonatal rat femur. NF 200-positive nerve processes (arrows), showing local enlargements (arrow heads), are running along bone trabaeculae (t) and

vessels (v). They are close to bone cells (Oc: osteoclast, Ob: osteoblast) and hematopoietic cells (H). Original magnification: X1000. (b) Deep

metaphysis of neonatal rat femur at the level of the afferent vessel. Tyrosine hydroxylase is present in these nerve processes (arrows) showing local

dilatations (arrow heads) in contact with hematopoietic cells (H). Oc: osteoclast. Original magnification: X1500. (c) Deep metaphysis of neonatal rat

femur. MAP2-positive nerve processes showing dilatations (arrow heads) in contact with hematopoietic cells (H) and osteoclasts (Oc). Original

magnification: X1000. (d) Diaphysis of 15-day-old rat tibia. NF200 is present in nerve endings (arrow heads) in contact with hematopoietic

cells (H). Original magnification: x1000, from Serre et al.(1999).

29

proliferation in vitro (Bongenhielm et al. 1995) The NP, ADM (adrenomedullin) is also

secreted by many cell types after LPS exposure and stimulates growth of odontoblast-

like MDPC-23 cells (Zudaire et al. 2006; So et al. 1996; Musson et al. 2010).Therefore

NP and NTGF released from non-neuronal cells of the dental pulp and bone may

stimulate both indirect and direct repair of mineralised tissue by activating the immune

and nervous systems as well as MSC.

NGF is previously the only NTGF to be investigated regarding direct effects on

bone repair and direct application was found to enhance bone regeneration when applied

in a fracture model and stimulated the differentiation of osteoblastic cells (Wang et al.

2006; Xian and Zhou 2009; Paschalia et al. 2011). NGF and its receptor, p75NTR

,

exhibit overlapping expression in polarising/differentiating odontoblasts during

odontogenesis and dentinogenesis (see Table 1.2, pg 36) (Maas and Bei 1997; Mitsiadis

et al. 1992, 1995; Fried 2000). Moreover, recent data also indicates that NGF may also

be an important ligand involved in tissue inflammation and repair of the dentine-pulp

complex (Woodnutt et al. 2000; Shiomi et al. 2000; O’Hara et al. 2009). Moreover,

postnatally NGF was up-regulated on pulp fibroblasts and odontoblasts within injured

teeth. Therefore similarly to a direct effect of NGF on osteoblasts, this molecule may

conceivably have a more direct function during dental repair by stimulating odontoblast-

like cell differentiation (Arany et al. 2009; Kurihara et al. 2007).

1.3.3 The role of GDNF in mineralised tissue repair

Sympathetic innervation has also been implicated in tooth development and

repair (Fried et al. 2000; Hildebrand 1995). Indeed the dental pulp is highly innervated

and contains sympathetic fibres of the trigeminal nerve that secrete NTGFs and

30

neuropeptides (Fried et al. 2001) (Figure 1.7, pg 31). Similarly to osteoblasts and

BMSC within bone, the nerve fibres within pulp are closely associated with

odontoblasts and can extend into the dentine (as described above) (Byers et al. 2003;

Egan et al. 1996) (Figure 1.6, pg 28, 1.7, pg 31 and 1.7.1, pg 32) and for animals of

lower evolutionary order such as polyphyodont teleost, (fish that continuously produce

teeth) nerve fibres have also been implicated in regulating odontogenesis (Hildebrand et

al.1995). Innervation of the tooth begins during the bud stage of development and this is

also at approximately the same time point when the mesenchyme expresses odontogenic

potential (Luukko et al. 1997). Due to this coordinated timing this has resulted in the

suggestion of a role for nerves in the regulation of odontogenesis (Luukko et al. 1996;

Hildebrand et al.1995). However, the location of GDNF and associated receptors

appears independent of the presence of the local nerve supply (Hellmich et al. 1996;

Luukko et al. 1996; Lukko et al. 1997), indicating that NTGF secretion within the

dental papilla may not necessarily emanate from the trigeminal nerve and may originate

from the non-neuronal DPC/MSC. Specifically, GDNF, GFRα1 and RET expression are

overlapping within sub-odontoblast and odontoblastic regions during late bell stage and

early postnatal days when the polarisation and differentiation of odontoblasts occurs

(Table 1.2, pg 35) suggesting a role for GDNF during odontogenesis and

dentinogenesis. Indeed NGF and GDNF have been suggested to direct the growth of the

trigeminal nerve into the dental papilla at the bud stage, however a causative effect of

GDNF or NGF has not been definitively demonstrated (Kvinnsland et al. 2004; Luukko

et al. 2008; Lillesaar et al. 2003). Moreover, the retrograde transport of NGF and

GDNF occurs away from the dental pulp into the main trigeminal nerve (Thoenen et al.

1988; Kvinnsland et al. 2004).

31

(i) (ii) (iii)

Figure 1.7 Innervation of the head and neck region and teeth. (i) The trigeminal nerve is the sensory nerve to the face also called cranial

nerve five (Cn V). It has three main, afferent branches from the face; known as Ophthalmic Va. Maxillary Vb and Mandibular Vc.The Vb and c

innervates the teeth (ii) Schematic showing gross anatomy of the teeth and entry of the nerve fibres into the dental pulp via the root canals (iii)

Trigeminal nerve density in peripheral pulp and dentine of adult rat molars is greatest near the tip of the pulp horn on one side (**) and is less

dense on the other side (*). The asymmetric density decreases towards mid-crown, and the innervation is missing from the dentine at the floor of

the crown. Nerve fibres were labelled by radioactive axonal transport and autoradiographic detection (modified from Byers and Kish, 1976;

Byers, 1984). Bars = 100 nm. From Byers et al. (2003).

32

Figure 1.7.1 Demonstration of nerve fibres in (A) human teeth using immunohistochemical staining for p75NTR

and (B) rat teeth with

immunohistochemical staining for Calbindin (CB). (A) The nerve fibres extend from the subodontoblastic nerve plexus (SN) towards the

odontoblast layer (OB). Arrows indicate the immunopositive Schwann cells. Reproduced from Maeda (1996), Bar = 100 nm. (B) Thin CB-

immunopositive nerve fibers penetrate into the predentin beyond the odontoblast cell layer (OB) (arrows) D: dentine. Bar = 100 nm, from

Miyawaki et al. (1997).

B A

33

GDNF is therefore detected during tooth development (Nosrat et al. 1998, 2002; Lukko et al.

1997; Fried et al. 2000) and up-regulated during repair in other tissues (Eberling et al. 2009;

Cheng et al. 2008). GDNF up-regulation from DPC or MSC may promote repair by modulating

nervous and immune systems and mineralised tissue responses. Neither GDNF nor other GFLs have

been studied in respect to their direct effects on the functioning within non-neuronal physiology of

mineralised tissues until recently (this study; Gale et al. 2011; 2012). Indirect evidence however

demonstrates that GDNF is likely to be important during tooth development. Indeed GDNF null

mice fail to form dentine and enamel, despite trigeminal nerve development being unaffected (De

vincente et al. 2002). Interestingly, Waardenburgs syndrome (resulting from Pax3/SOX10

mutations), has a cross over in definition and symptoms with Hirschsprung’s disease where ~50%

of Hirschsprung cases have inactivating mutations of RET (Tassabehji et al. 1993; Wang et al.

2010b; Yang et al. 2012; Bondurand et al. 2007; Pelet et al. 1998) and phenotypes of both diseases

may present with missing teeth and defects in craniofacial and limb bones (Eigelshoven et al.

2009; Bandyopadhyay et al. 1999; Zelzer and Olson 2003; Pingault et al.1998; Parisi 2002). These

data suggest that GDNF-RET expression within the developing dental papilla may reflect the

importance of this signalling complex in neural crest derived cells during odontogenesis,

dentinogenesis, craniofacial and long bone development.

MGIF (murine glial cell-derived neurotrophic factor inducible transcription factor) is

homologous to TIEG (TGF-beta-inducible early-response gene) (Yajima et al. 1997; Subramaniam

et al. 2005; Consales et al. 2007; Hefferan et al. 2000) and is implicated in osteoblast

differentiation. Furthermore upstream of GDNF intracellular signalling, the Lef-1 transcription

factor, is reported to stimulate odontoblast and osteoblast differentiation (Yokose and Naka 2010;

Hoeppner et al. 2011; Miletich and Sharpe 2003) which has a putative consensus site for binding to

34

the GDNF promoter (Tanwar et al. 2010). Taken together, these data further suggest that GDNF

may play a role in osteoblast and odontoblast differentiation during development and repair.

1.4. Aims of this study

Hypothesis: GDNF affects non-neuronal/mesenchymal dental pulp cell (DPC) behaviour and

may promote the regenerative responses in mineralised tissues.

The aims of the study were to investigate the expression and action of GDNF within in vitro culture

models of mesenchymal cells capable of forming mineralised tissues. For this purpose, DPC, BMSC

and cranial osteoblast-like cells were used.

Research objectives

a) To identify suitable DPC culture models and compare with BMSC and osteoblast-like cultures to

allow the study of GDNF action in vitro.

b) To study cellular proliferation, differentiation and in vitro mineralisation in response to GDNF in

vitro.

c) To assess the effects of GDNF under conditions reflecting some aspects of the inflammation and

infection processes that occur during disease and repair.

35

STAGES OF TOOTH DEVELOPMENT

Transcript

detected

Bud stage Cap stage

Bell stage Postnatal/neonatal Postnatal

References

GDNF Facial cartilages;

Jaw mesenchyme

Dental epithelium

Jaw, mesenchyme,

Dental papilla; Dental

follicle; Dental

epithelium

Dental papilla;Pre-

odontoblasts

Pre-odontoblasts, odontoblasts,

Sub-odontoblastic mesenchyme;

Sub-odontoblastic layer

Odontoblast and

Sub-odontoblast

region

(Nosrat et al. 2002;

Luukko et al. 1997;

Nosrat et al. 1998;

Fried et al. 2000)

RET Dental

mesenchyme

Dental papilla

mesenchyme

Dental papilla

mesenchyme

Sub-odontoblastic region and

odontoblastic region

Not reported or

detected

(Luukko et al. 1997;

Nosrat et al. 1997)

GFRα1 Not reported or

detected

Dental and outer

epithelium

Dental and outer

epithelium;Pre-

odontoblasts and sub-

odontoblastic region

Odontoblasts and Sub-

odontoblast region

Pre-odontoblast region

Not reported or

detected

(Luukko et al. 1997;

Nosrat et al. 1997)

Table 1.2 GDNF, RET, GFRα1, NGF and p75NTR transcripts detected within the developing tooth. The data cited above was based on different in vivo

models; Luukko et al. (1997) and Nosrat et al. (1997) used murine, Nosrat et al. (2002) used human and Hildebrand et al. (1995), Byers et al. (1990),

Mitsiadis et al. (1992) used rat dental tissues.

36

STAGES OF TOOTH DEVELOPMENT

Transcript

detected

Bud stage Cap stage

Bell stage

Postnatal/neonatal Postnatal

References

NGF Not reported or

detected

Dental follicle,

outer epithelium

Pre ameloblast,

Pre odontoblasts

Not reported or detected Not reported or

detected

(Hilldebrand et

al. 1995;

Mitsiadis et al.

1992)

P75NTR

Dental

epithelium

Enamel

epithelium,

Dental papilla

(transient

expression)

Dental papilla

(transient

expression)/pre

odontoblasts

Sub odontoblastic Sub-

odontoblastic

(Byers et al.

1990; Hildebrand

et al. 1995;

Mitsiadis et al.

1992)

Table 1.2 continued GDNF, RET, GFRα1, NGF and p75NTR mRNA detection within the developing tooth. The data cited above was based on different

in vivo models; Luukko et al. (1997) and Nosrat et al. (1997) used murine, Nosrat et al. (2002) used human and Hildebrand et al. (1995), Byers et al. (1990),

Mitsiadis et al. (1992) used rat dental tissues.

37

CHAPTER 2 MATERIALS AND METHODS

2.1 Cell isolation

2.1.1 Culture of bone marrow stromal/stem cells (BMSC)

Femurs of male Wistar rats (Charles River Laboratory, UK) aged 4-6 weeks old

were dissected out and the muscle stripped from the bones using a scalpel, scissors and

forceps. The femurs were then placed into universal tubes containing medium

consisting of αMEM (Biosera) supplemented with 10 % FBS (Fetal Bovine Serum),

penicillin (100 µg/ml), streptomycin (100 µg/ml) and (2.5 µg/ml Amphotericin B

(Sigma-Aldrich). The femurs were sprayed with 70 % ethanol and the ends (epiphyses)

were removed using a scalpel blade and were then flushed through with 10 ml medium

using a needle (~22gauge) and 20 ml syringe (BD sciences). The bone marrow was

collected into a 50 ml centrifuge tube (Falcon). Cells were collected by centrifugation

(5804R, Eppendorf) at 800 rpm for 5 mins and pellets were re-suspended in 5 ml of 10

% FBS supplemented αMEM medium. Cell suspensions were seeded into 75 cm2

flasks

(Nunc) containing 10 % FBS supplemented αMEM (Biosera).

2.1.2 Dental pulp extraction

Incisors (upper and lower) were dissected from 4-6 week old male Wistar rats. The

extracted teeth were briefly submerged in 70% ethanol for disinfection and then

immediately placed into universal tubes containing 20% FBS / αMEM (Biosera)

supplemented with penicillin (100 µg/ml) streptomycin (100 µg/ml) and (2.5 µg/ml)

Amphotericin B (Sigma-Aldrich). Teeth were placed onto a glass slide within a petri

38

dish and the pulp removed from the tooth using forceps and a scalpel (Figure 2A).

Medium was added to the pulp tissue to prevent it from becoming desiccated.

2.1.3 Explant-derived dental pulp cells (DPC)

Pulpal tissue was placed onto a sterile glass slide and chopped coarsely to ~

2mm2 pieces using a scalpel, prior to transfer to culture flasks containing 1 ml 20 %

FBS supplemented αMEM (Figure 2B, pg 42). These explants were gently pressed onto

the bottom of the flask using the flat side of the scalpel blade. Cells were incubated

overnight in a humidified atmosphere at 37oC with 5 % CO2 and the following day a

further 4 ml of 20 % FBS supplemented αMEM was added to the culture. Medium was

changed every 3 days until cultures reached 80 - 90 % confluence.

2.1.4 Enzymatically-derived DPC

Extracted pulp was minced (see Figure 2A, pg 42) using a scalpel and then

transferred to a 15 ml centrifuge tube (Falcon) containing 5 ml trypsin [2.5g/l of trypsin

in 0.38g/l of EDTA (Invitrogen)] and digested for 30 mins at 37oC. Tissue samples were

agitated every 5 mins using a 1 ml Gilson pipette tip to aid dissociation, then after 30

mins, 5 ml 10 % FBS supplemented αMEM was added to halt the action of the trypsin.

Subsequently the cell suspension was passed through a 70 µm filter (Millipore) and

centrifuged for 5 min at 800 rpm. The supernatant was removed using vacuum

aspiration and the cell pellet re-suspended in 1 ml of αMEM and seeded into a 25 cm2

flask (Nunc). Cells were incubated overnight humidified at 37oC, 5% CO2 to enable

attachment and the following day a further 4 ml of 20% FBS supplemented αMEM was

added to the culture. Medium was changed every 3 days until cultures reached 80 – 90

% confluence.

39

A

B

Figure 2 (A) Dental pulp was extracted from the tooth. (B) Dental pulp was minced

using a scalpel blade prior to enzymatic digestion.

Extracted

dental

pulp on

glass slide

Minced dental

pulp

Mineralised

tooth with

dental pulp

removed

Extracted

dental

pulp on

glass slide

Mineralised

tooth with

dental pulp

removed

40

2.1.5 Cell lines culture

A C6 glioma cell line of isolated rat glial cells derived from a chemically

induced glial tumour by Benda et al. (1968), a mouse MDPC-23 cell line isolated from

fetal mouse dental papillae (Hanks et al. 1998) and the mouse osteoblast MC3T3-E1

cell line established from neonatal calvarial bone of C57BL/6 mice (Wang et al. 1999)

were established from cryopreserved stocks (original sources of cell lines were ATCC

except MDPC-23 cells which were kindly donated by Jacques Nör’s Laboratory,

University of Michigan, US).

2.1.6 Cell culture

Explant-derived and enzymatically-derived DPC were cultured as above (2.1.3

and 2.1.4), MC3T3-E1 cells and BMSC were cultured using the same reagents except

10% FBS (Biosera) was used. MDPC-23 and C6 glioma cells were cultured using

DMEM (Biosera); 10% FBS, Penicillin (100 µ/ml) streptomycin (100 µg/ml), L-

glutamine (2 mM) (Sigma-Aldrich). All cells were cultured in either 25 cm2 (primary

cells, passage 0-1) or 75 cm2 flasks (Falcon). Cells were detached upon reaching 80-90

% confluence using trypsin/EDTA. Cell suspensions were transferred to tubes (Falcon)

size 15 ml or 50 ml respectively, depending on flask size and centrifuged at 800 rpm for

5 mins to pellet the cells. Supernatants were removed by using a vacuum pump and

cells re-suspended in 1 ml of αMEM for counting or subculture. DPC and BMSC were

expanded and used at passage 2-4 for subsequent experiments.

41

2.1.7 Cell counting

A 100 µl sample of cell suspension was mixed with 100 µl Trypan blue (Sigma); the

unstained viable cells were counted using a Neubauer haemocytometer slide under a

variable relief contrast (VAREL) microscope (Zeiss Axiovert 25). A minimum of three

counts per sample were obtained and average cell number was calculated. Cell

suspensions were then diluted accordingly and seeded at densities and volumes

presented in Table 2, pg 45- 46.

2.1.8 Preparation of serum-free medium

Bovine serum albumin (BSA) supports cell cultures by acting as a buffer,

antioxidant and carrier for proteins, lipids and minerals (He and Carter 1992; Nicholson

et al. 2000). BSA (fraction V, Sigma) was reconstituted with dH2O to a 1 % solution

(w/v), then heat inactivated at 56oC for 30 mins. This solution was filter sterilised using

a 0.2 µm filter (Millipore) and stored at -20 oC prior to use. Serum-free medium

consisted of 0.1 % BSA in αMEM, penicillin (100 µg/ml) streptomycin (100 µg/ml) and

Amphotericin B (2.5 µg/ml) (Sigma-Aldrich). After seeding into an appropriate tissue

culture vessel the cells were incubated in a humidified 5% CO2 incubator at 37oC.

2.2 Preparation and storage of GDNF

Human GDNF is highly homologous to mouse or rat GDNF (Wang et al. 1998).

rh GDNF [recombinant human GDNF produced in E.coli (Amgen) ] was supplied at a

concentration of 10 mg/ml and aliquots of 1 µg/ml in 0.1 % BSA/αMEM were stored at

-80oC. On thawing GDNF was diluted with medium to the relevant concentrations used

for the in vitro experiments. Before adding GDNF to the cultures the medium was

42

replaced with freshly prepared medium, containing either 10 % FBS or 0.1 % BSA

supplemented αMEM (serum-free medium). Cells were then cultured for 48 hours,

except for the mineralisation assays where cells were cultured for 4 days to obtain

confluence before osteogenic conditions were applied. Cells were incubated in a

humidified 5 % CO2 incubator at 37oC, for periods shown in Table 2, pg 45- 46.

A

B

Figure 2.1 Phase-contrast images of (A) enzymatically-derived and (B) explant-

derived DPC.

Explanted

pulpal

tissue

43

2.2.1Preparation and storage of tumour necrosis factor alpha (TNFα)

Human, rat and mouse TNFα are highly homologous, the sequences display 80-

89% homogeneity (Kwon et al. 1993; Marmenout et al. 1985). rh TNFα [ produced in

E.coli (Peprotech)] was centrifuged and reconstituted using 0.1 % BSA/αMEM. 1 µg/ml

aliquots were stored at -80oC. On thawing, TNFα was serially diluted using serum-free

or serum supplemented medium with or without GDNF (100 ng/ml). The same protocol

was used as for the GDNF experimental analysis (section 2.2; see Table 2, pg 45-46).

2.2.2 Preparation and storage of nerve growth factor beta (NGF-β)

Mouse, rat and human NGF are highly homologous (Hallböök et al. 1991;

Ullrich et al. 1983). NGF has three subunits, alpha, beta and gamma (Baker et al.

1974). It is the beta subunit that exerts the majority of the biological effects of NGF

(Shooter 2001). rh NGF-β [ produced in E.coli (Peprotech)] was reconstituted at 100

µg/ml in 0.1 % BSA/αMEM and 1 µg/ml aliquots stored at -80oC. On thawing NGF was

diluted in either serum-free or serum supplemented medium and the same protocol was

used as for the GDNF experiments (section 2.2; see Table 2, pg 45-46).

2.2.3 WST-1 viable cell number assay

To determine the number of viable cells the WST-1 assay (Roche) was

performed. This assay was originally developed by Ishiyama et al. (1993) (see also

Scheven et al. 2009). In accordance with the manufacturers guidelines 10 µl of WST-1

reagent per 100 µl of medium was incubated with cell cultures for 60 mins at 37oC. In

viable, metabolic active cells, WST-1 is converted to form a coloured soluble formazan

dye by dehydrogenases. A universal plate reading spectrophotometer (ELx800, Biotek)

44

was used to quantify the amount of formazan dye formed at 450/630nm. The 630 nm

wavelength was used as a reference wavelength, this represents background readings

that was subtracted from the test spectrophotometry reading. The linear relationship

between the formazan dye product and seeding density (up to 50,000 cells per well)

under differing serum conditions was subsequently used to determined the number of

viable cells under different experimental conditions.

2.2.4 RET receptor inhibition (RPI-1)

RPI-1 (1, 3-Dihydro-5, 6-dimethoxy-3-(4-hydroxyphenol) methylene)-2H-indol-

2-one) (Merck Chemicals Ltd) was used to inhibit RET. RPI-1 was reconstituted to 5

mg/ml in DMSO, subsequently aliquots at 300 µM/0.2 % DMSO/ αMEM were stored at

-20oC prior to use. The RPI-1 inhibitor was serially diluted in serum-free medium with

or without GDNF (100 ng/ml) and incubated for a further 48 hours with cell cultures.

2.2.5 GFRα1 receptor inhibitor (PI-PLC)

PI-PLC hydrolyses the extracellular portion of the GFRα receptor (and other

proteins that are GPI-linked glycosyl-phosphatidylinositol) from the membrane surface

(Low 1989). Thus PI-PLC cleaves phosphatidylinositol and this removes the

extracellular receptor protein and hence the binding site for GDNF. PI-PLC (Sigma)

62.5 units/ml was stored at 2-8oC. Treatment of the cells was started 1 hour prior to

addition of GDNF (to allow adequate time for cleavage of the receptor from the cell

membrane). The medium was then removed and replaced with respective concentrations

of PI-PLC (as above) in serum-free medium with or without GDNF (100 ng/ml) and

incubated for a further 48 hours with cell cultures (see Table 2, pg 45- 46).

45

Table 2 Summary of experimental assays and conditions used.

Assay Applied Experiment/Analysis Culture ware and seeding

density used

Duration/time

point examined

Detection method Additional reagents

prepared

WST-1 viable

cell number

Standard curve

determination

96 well plates; 5x103

cells in

200 µl per well

30 mins, 24 and 48

hours

Spectrophotometry at

450/630 nm

0.1 % BSA

WST-1 cell

viability

Comparison of serum

conditions all cell types


cells in

200 µl per well

48 hours, 3, 5 10

days

Spectrophotometry at

450/630 nm

0.1 % BSA

WST-1 cell

viability

Effect of GDNF

On all cell types


cells in

200 µl per well

48 hours Spectrophotometry at

450/630 nm

GDNF

0.1 % BSA

WST-1 cell

viability

Effect of S.mutans on DPC

and MC3T3-E1 cells


cells in

200 µl per well


450/630 nm

S.mutans

WST-1 cell

viability

Effect of TNFα on DPC and

MC3T3-E1 cells


cells in

200 µl per well


450/630 nm

TNFα; GDNF; 0.1 %

BSA

WST-1 cell

viability

Effect of NGF on DPC 96 well plates; 5x103

cells in

200 µl per well


450/630 nm

NGF; 0.1 % BSA

WST-1 cell

viability

Inhibition of RET and

cleavage of GFRα receptors


cells in

200 µl per well


450/630 nm

PI-PLC; GDNF; 0.1 %

BSA; RPI-1

46

Assay Experiment Seeding density Duration/time

point

Detection Additional reagents

prepared

ALP ALP levels for all cell types 96 well plates; 5x103

cells in 200

µl/ well


405/630 nm

Buffer for pNPP tablets

BrdU Proliferation assay for DPC and

MC3T3-E1 cells


cells in

1ml/well

24 and 48 hours Light microscopy. Cell

counts calculated using

image J cell counter add

in.

Buffers; fixative;

antibodies; GDNF

Live dead Number of live and dead cells

determined


cells in

1ml/well

24 and 48 hours Fluorescent

microscopy. Cell counts

BrdU; Acridine orange; 0.1

%BSA; ethanol fixative

Caspase-3/-7 Levels of apoptosis determined

for DPC


cells in 200

µl/well

48 hours Luminometer 0.1 % BSA.

LDH Levels of lactate

dehydrogenases (LDH)

determined the levels of cell

death for all cell types


cells in 100

µl/ well


490/630 nm

0.1 % BSA. Kit preparation

Mineralisation Levels of calcified ECM for

MDPC-23, DPC and MC3T3-

E1 cells

48 well plates; 2.5x104 cells at

200 µl/well. 35mm dishes; 5X104

7, 24 and 48 days Spectrophotometry

450/630 nm

Alizarin Red 2 % solution.

Ammonium hydroxide; 0.1

% BSA

Table 2 continued. Summary of experimental assays and conditions used.

47

2.2.6 BrdU (5-Bromo-2’-deoxy-uridine) assay

Cells were cultured within multiwell plates (see Table 2, pg 45- 46) and after 24

hours medium was replaced with serum-free or serum supplemented medium with or

without GDNF (100 ng/ml). After a further 24 or 48 hours the BrdU assay was

performed using the 5-Bromo-2’-deoxy-uridine labelling and detection kit II (Roche)

according to the manufacturer’s guidelines. Cells were grown in 12-well tissue culture

plates, incubated with the DNA-intercalating BrdU for 60 mins. The cells were then

fixed and fractured to later allow antibody access to the BrdU labelled DNA. The

fixation procedure included fixing cells with 70 % ethanol for 20 mins at room

temperature and followed by fixation using 50 mM glycine/70 % ethanol at pH 2.0 for

20 mins. Finally cells were fractured by incubating at - 20oC for 20 mins. The DNA

labelled 5-Bromo-2’-deoxy-uridine was detected using antibodies and visualised by

ALP conjugated secondary antibody reacted with BCIP (5-bromo-4-chloro-3-indolyl

phosphate, toluidinium in dimethylformamide). Mayer’s haematoxylin was used as a

counterstain and a variable relief contrast Zeiss Axiovert 25 microscope was used to

observe BrdU labelled nuclei that appeared brown/black. BrdU stained and non-stained

nuclei were counted in at least nine fields of view for each test condition.

2.2.7 Live and dead assay

Cultures were washed with 500 µl PBS and then ethidium bromide (EthBr) and

flurochrome acridine orange (AO) were added (4 µM in PBS) for 5 mins. Dead cell

controls were fixed cells using 70 % ethanol/PBS and all fixed (dead) cells appeared

orange. EthBr crosses damaged cell membranes and intercalates with DNA and emits

48

fluorescent light appearing orange. AO binds to nuclear DNA and all live cells take up

the AO stain across the cell membrane and nuclei appear green however dead cells with

ruptured/leaky membranes take up EthBr and over stain the green AO to appear orange.

Cells were visualised and counted using a Nikon Eclipse fluorescent microscope with

480-520 nm filter (530 nm max emission for AO and 590 nm for EthBr). For each

experiment three wells were counted per test condition and within each well three fields

of view were counted and numbers of live/dead cells were recorded.

2.2.8 Lactate Dehydrogenase (LDH) assay

The LDH Cytotoxicity Detection Kit (Roche) was utilised according to the

manufacturer’s instructions. After 48 hours samples of cultured cells were exposed to

the supplied lysis buffer for 20 mins to act as a positive control. 100 µl of supernatant

was removed and incubated with the manufacturers’ catalyst and dye reaction mixture

for 30 mins at room temperature (RT). After 30 mins the stop reaction mixture was

added at 25 µl per well and samples were placed on an orbital plate shaker for 20

seconds. The LDH levels released from damaged cell membranes were then quantified

using a multiwell plate reading spectrophotometer (ELx800Biotech) at a wavelength of

490/630 nm.

2.2.9 Caspase Glo 3/7 assay

After 24 hours culture in white walled multiwell plates (Microlite; 7567

Thermoscientific) the medium was replaced and serum-free, serum supplemented and

serum-free medium containing GDNF (100 ng/ml) was added to the cultures for a

further 48 hours. The Caspase Glo 3/7 assay (Promega) was performed according to the

49

manufacturer’s instructions. Caspases cleave the caspase specific DEVD amino acid

sequence within the supplied luminogenic substrate to form a substrate for the luciferase

enzyme that is luminescent following cleavage. Luminescence from the reaction was

measured by taking continuous readings (0-120 mins) using a multiwell plate

luminometer (LB96v Bethold Technologies) and data from the time point of maximum

peak emission used for analysis. Plates were incubated within the luminometer at 22oC.

2.3 Streptococcus mutans (S.mutans) analysis

Heat inactivated S.mutans [(Streptococcus mutans Clarke 1924 AL) original

source from the ATCC No. 25175] were stored at -80oC at 7.55 x 10

9/ml prior to use.

On thawing the bacterial suspension was diluted to 5x106 S.mutans/ml and further

serially diluted to obtain a ratio of 1000, 100, 10 and 1 S.mutans per DPC or MC3T3-E1

cell. DPC or MC3T3-E1 cell cultures were seeded at 5x103 per well prior to addition of

bacteria in serum supplemented medium with or without 100ng/ml GDNF. After

incubation for 48 hours the WST-1 assay was performed to quantify viable cells using a

plate reader (ELx800 Biotek).

2.3.1 Mineralisation assay

Cells were seeded into 35 mm dishes under standard culture conditions and

medium. After 4 days medium was changed to include osteogenic supplements which

consisted of 10-7

M dexamethasone (DEX) 50 µg/ml, L-ascorbic acid, 10 mM β-

glycerophosphate (Sigma) (Jaiswal et al. 1997). This osteogenic medium with or

without 100 ng/ml GDNF was freshly prepared prior to use and changed every other

day for the duration of the experiments for up to 28 days.

50

For serum-free mineralisation assays the cells were first seeded into 48 well

plates under standard serum conditions for 24 hours. After 24 hours the medium was

changed to serum-free medium containing 0.1 % BSA, osteogenic supplements with or

without GDNF as described above. L- glutamine is essential for the survival and growth

of fibroblasts under low serum conditions (Eagle et al. 1955) therefore an additional,

2mM L-glutamine was added to the osteogenic medium. After 4 weeks culture under

serum-free osteogenic conditions cultures were stained with Trypan blue to assess the

levels of viable cells under the experimental conditions. Culture medium was aspirated

and 50 µl Trypan blue: 50 µl PBS was added to each well and viewed immediately. The

staining was visualised using an inverted microscope (Zeiss Axiovert 25).

Figure 2.2 Trypan blue staining of DPC in (A) serum-free cultures and (B) serum

supplemented cultures after 4 weeks culture with osteogenic supplements. Red

arrow shows dead cell identified with Trypan blue staining and blue arrows indicate live

cells.

51

Alizarin red stain (ARS) forms salts with calcium, replacing the anions in

calcium (Puchtler et al. 1969). The pH of the ARS stain was adjusted to pH 4.1 using

ammonium hydroxide. ARS was made freshly to ensure pH 4.1 was maintained due to

the solubility of calcium deposits at pH 4.5-5 (Puchtler et al. 1969). Cultures were

washed with 200 µl PBS per well, then fixed with 10 % formalin for 10 mins. Formalin

was aspirated. 200 µl of 2 % ARS was added to the cultures at room temperature (RT)

and agitated on an orbital plate shaker for 20 mins. The unincorporated ARS was

aspirated and the cells were washed gently with PBS, then with dH2O for 5 mins. The

dH2O was drained by inverting the plate/dish.

For the post staining of 35 mm dishes and 48-well plates, 800 µl and 400 µl of

acetic acid (10 %) was added, respectively and incubated for 30 mins RT on an orbital

plate shaker. Dishes and wells were scraped and contents placed into centrifuge tubes.

Tubes were mixed using a vortex for 30 seconds, heated at 85oC for 10 minutes and

after 5 mins cooling on ice, centrifuged at 1500 rpm for 15 minutes. Then 150 µl (for

dishes) or 100 µl (per well) of 10 % ammonium hydroxide was added to each sample to

neutralise the acid and 150 µl from each sample was added to multiwell plates in

triplicate. The concentration of ARS staining per eluted sample was determined against

a standard curve using a plate reader at 405/630nm (ELx800, Biotek).

52

Figure 2.3 Linear standard curve for ARS. Results expressed as average absorbance

unit ( n=2).

2.3.2 Alkaline phosphatase (ALP) assay

Cells were rinsed using 100 µl PBS per well of multiwell plates (see Table 2.0),

then lysed using 0.1 % triton X100. pNPP (p-Nitrophenyl Phosphate) tablets were

added to DEA buffer to make 1 mg/ml pNPP and incubated at RT for 10 mins. The

DEA buffer contains Mg2+

and Zn2+

ions enabling optimum activity of alkaline

phosphatase. The DEA buffer stock at 10X concentration consisted of 500 mg MgCl2,

400 mls dH2O and 48.5 ml diethanolamine, pH was adjusted to pH 9.8 using

concentrated NaOH or HCl. pNPP is hydrolysed by ALP to the yellow coloured p-

nitrophenol and its levels were determined using a multiwell plate reader at 405/630

nm.

53

2.4 Reverse transcriptase polymerase chain reaction (RT-PCR) analysis

Gene expression was investigated using semi-quantitative RT-PCR (sqRT-PCR)

analysis. RNA isolation was performed using the RNeasy mini-kit (Qiagen) with DNase

treatment according to the manufacturer’s instructions. Dissected tissue samples were

initially disrupted using an Ultraturrax homogeniser (Netzgerat T8.01 aboratiechnik)

within 700 µl of lysis buffer (10 µl β-Mercaptoethanol per 1ml buffer RLT). 700 µl

lysis buffer was added per confluent (80 - 90 %) cultures in 75 cm2 flasks followed by

vigorous pipetting to detach and disperse the cells. After ~ 5 mins 70 % ethanol was

added to all samples and two aliquots per sample of the total volume (1400 µl) pipetted

into RNeasy mini- column sequentially (the total column volume per aliquot does not

exceed 700 µl) and centrifuged for 15 seconds at 10,000 rpm. The flow through was

discarded after each centrifugation. All centrifugation steps were performed using an

Eppendorf 5415 D centrifuge (Eppendorf, UK). Contaminating DNA was removed by

performing the DNase digestion steps; 350 µl RW1 manufacturers buffer was added per

column, centrifuged at 10,000 rpm for 15 secs and flow through discarded. 15 µl DNase

I (inhibitor) and 60 µl RDD manufacturer’s buffer was added per column and incubated

at RT for 15 mins. Next 350 µl RW1 buffer added to each column and centrifuged at

10,000 rpm for 15 secs, then flow through was discarded. Subsequently, 500 µl RPE

buffer was added to each column within a new collecting tube and centrifuged at 10,000

rpm for 15 secs, then flow through was discarded. This step was repeated except

centrifugation took place for 2 mins then columns were placed into new collecting tubes

and centrifuged for a further 1 min at full speed. The RNA was eluted by placing the

column into an Eppendorf and adding 30 µl RNase free water. RNA was collected by

54

centrifugation for 1 min and the collected RNA placed onto ice whilst purity and

quantity were assessed. Purity and quality of the RNA was determined by measuring

absorbance values at 260 nm (A260) and 280 nm (A280) using a Biophotometer

(Eppendorf, UK) and by visualisation on agarose gels (see section 2.4.1, pg 55). An

absorbance ratio of between 1.8 to 2.1 and distinct double bands (18S and 28S) of RNA

observed on gels signified high purity and quality of the isolated RNA. RNA was

stored at - 80oC prior to use.

RNA was reversed transcribed using the Omniscript Reverse Transcription kit

(Qiagen) according to the manufacturer’s instructions. 50 ng - 1 µg RNA was incubated

with 2 µl sterile molecular grade water (Merck) at 65oC

for 5 min to denature RNA

secondary structure and quenched on ice prior to use. A master mix containing 1X RT

buffer, 0.5 mM dNTPS, 1 µM Oligo-dT primer, 0.5 u/µl RNase inhibitor, 0.2 u/µl

omniscript reverse transcriptase were combined and added to the RNA solution making

a total volume of 20 µl per sample. This reaction mix was incubated in a Mastercycler

gradient thermal cycler (Eppendorf, UK) for 60 mins at 37oC and then 95

oC for 5 mins.

YM-30 Microcon centrifugal filter devices (Millipore) were used to purify and

concentrate resultant cDNA. 20 µl of cDNA in solution was added to 500 µl in

molecular biology grade water, transferred to YM-30 microcon assemblies and

centrifuged (Centrifuge 5415D, Eppendorf) for 6.5 minutes at 10,000 rpm (Centrifuge

5415D, Eppendorf). The eluate was discarded and this process was repeated until

approximately 30 μl cDNA in solution remained on the surface membrane of the filter.

The microcon centrifugal filter device was inverted in an Eppendorf tube and

centrifuged at 1000 rpm (Centrifuge 5415D, Eppendorf) for 3 mins to collect the cDNA.

55

The purity and concentration of the cDNA were analysed by determining absorbance

readings at 260 nm and gel electrophoresis (see section 2.4.1). cDNA was stored at -

20oC prior to use.

50 ng of the cDNA, primers (25 mM) (see Table 1.0) and

REDTaq®ReadyMix™ PCR Mix (with MgCl2) were gently combined and transferred

to a Mastercycler gradient thermal cycler (Eppendorf, UK) and amplified using repeated

cycles. Reactions were amplified following an initial denaturation step of 5 min at 94

°C, a typical amplification cycle consisted of denaturing 94 °C for 20s, annealing

(temperatures shown in Table 2.1, pg 58) for 20 s and extension at 68 °C for 20 s ending

with a 10 min extension at 72 °C. Following the designated number of cycles (Table

2.1), 6 μl of the reaction mix was removed. Primers (Table 2.1, pg 58) were designed

from NCBI mRNA sequences Primer-3 Primer design software

(http://biotools.umassmed.edu/bioapps/primer3_www.cgi). The PCR products were

analysed by agarose gel electrophoresis (see below section 2.4.1).

2.4.1 Agarose gel electrophoresis and image analysis

Non-denaturing agarose gels (1 % w/v for RNA and 1.5 % w/v for PCR) were

generated from lyophilised agarose powder (Helena Biosciences, UK) dissolved in 1X

Tris acetate EDTA (TAE) buffer, pH ~8.3, (Helena Biosciences UK). Mixtures were

heated in a microwave (Samsung, TDS) and the solution was poured into gel casting

trays. Ethidium bromide 50 µg/ml (Sigma) or 1X SYBR gold (Molecular probes)

nucleic acid stains along with combs were added to enable nucleic acid visualisation

and to form sample-loading wells prior to gel solidification at room-temperature.

56

Samples of 6 µl per well were added and separated by gel electrophoresis at 50-100V

in agarose gels submerged under 1x TAE (Tris-Acetate EDTA) running buffer with the

Hyperladder IV (Bioline, UK) as a nucleic acid standard.

Images of RNA and PCR products were visualised by UV illumination and

captured using a G:BOX system with the gene tools software (Syngene) used to perform

the densitometric analysis. Density values were normalised to glyceraldehyde-3-

phosphate dehydrogenase (GAPDH) housekeeping gene (Robbins and McKinney 1992)

control value by dividing target gene sample volume density by respective GAPDH

volume density.

2.5 Immunocytochemistry

Cells were seeded onto multispot slides (Hendley) at a density of 1.5 x104

Glioma cells, 2 x104 DPC, 2 x10

4 MC3T3-E1 cells, or 2.5 x10

4 BMSC and incubated

for 24 hours in standard serum conditions. Cells were then gently washed with warm

(37oC) PBS/1% BSA, fixed with cold acetone at - 20

oC for 5 mins, followed by

quenching of endogenous peroxidase using 3 % H2O2/PBS for 30 minutes. After

washing with PBS/1 % BSA, the slides were ―blocked‖ with 20 % goat serum/1 %

BSA/PBS for 1 hour to prevent non-specific antibody binding. Primary antibodies

against Gfrα1 (sc10716, polyclonal rabbit; SantaCruz) or against RET (sc167,

polyclonal rabbit; Santa Cruz) at 2 µg/ml/1 % BSA/PBS and controls of normal rabbit

serum (Sigma) 14 µg/ml/1 % BSA/PBS or 1 % BSA/PBS were added and incubated for

1 hour at RT. The substitution of 1 % BSA/PBS in place of primary antibody was used

to determine the specificity of the secondary antibody (shown in Figure 2.4 pg 59).

57

Biotinylated secondary antibody (link) was added at 1/200 dilution in 1 % PBS/BSA

and incubated for 1 hour at RT. Next streptavidin-HRP (label) was added 1/200 dilution

and incubated for 30 minutes at RT. The SuperSensitive™ Link-Label IHC Detection

System (Biogenex LP000-UL) was used to detect and visualise the antibody labelling.

DAB solution (0.01g 3,3-diaminobenzidine tetrahydrochloride plus 25 µl H2O2 in 20

mls PBS) was added until a golden brown colour developed, subsequently, the slides

were counterstained with Mayers haematoxylin for 5-15 mins. Cells were then

dehydrated through sequential steps in 100 %, 95 % then 70 % alcohol and placed into

xylene and mounted using XAM (BDH Laboratory). Stained cells were examined under

a variable relief contrast (VAREL) microscope (Zeiss Axiovert 25)

2.6 Statistical analysis

Statistical and data analyses were performed using Microsoft Excel and Minitab

(version 15 and 16). Statistical analyses comparing groups were undertaken using the

Student’s t test and in the case of more than two groups regression/ANOVA with

Tukeys posthoc test. A P-value of < 0.05 was considered to be significant.

58

Gene Primer sequence (F-forward; R-reverse) 5’ 3’

Tm Accession no. Product Cycle No.

GAPDH F-CCCATCACCATCTTCCAGGAGC;R-CCAGTGAGCTTCCCGTTCAGC

60 NM_017008 473bp

21-27

GDNF F-GAGGAATCGGCAGGCTGCAGCTG;

R-GATACATCCACATCGTTTAGCGG

60 NM_019139

364bp 33-38

rRET F-TCAGGCATTTTGCAGCTATG;

R-TGCAAAGGATGTGAAAGCAG

62.5

NM_001110099.1 393bp 35-45

Gfrα1 F-AATGCAATTCAAGCCTTTGG;

R-TGTGTGCTACCCGACACATT

60

NM_012959.1 218bp 33-43

P75NTR

(NGFR)

F-CAAAGGACGGATTTCCTGAG;

R-AGCTCCTGGGGAGGAAAATA

60

NM_012610 323bp 37

CD105 F-TTCAGCTTTCTCCTCCGTGT;

R-TGTGGTTGGTACTGCTGCTC

60 NM_001010968 325bp 41-45

CD44 F-TGGGTTTACCCAGCTGAATC;

R-CTTGCGAAAGCATCAACAAA

60 NM_012924.2 392bp 33-37

PCNA F-TTGGAATCCCAGAACAGGAG;

R-CGATCTTGGGAGCCAAATAA

60 NM_022381.2 390bp 30-33

p27kip1 F-ACGGTTCCCCGAATGCTGGC;

R-CCCCCACCCAAGTTGCTTCTCT

60 NM_031762.3 483bp 30-42

IL-6 F-TGTGCAATGGCAATTCTGAT;

R-GAGCATTGGAAGTTGGGGTA

60 NM_012589 312bp 33

TNFα F-TCCGGCGGTGTCTGTGCCT;

R- CGGGGCAGCCTTGTCCCTTG

60 NM_012675 369bp 35

BSP F-ATGGAGATGGCGATAGTTCG;

R-TCCACTTCTGCTTCTTCGTTC

60 NM_012587.2 307bp 34-35

Cfbα1 F-GCCGGGAATGATGAGAACTA;

R-GGACCGTCCACTGTCACTTT

60 NM_053470.2 200bp 35

Table 2.1 Details of primers and assay conditions for PCR analysis. Tm indicates

annealing temperature oC, rRET, rat RET primer sequences.

59

Figure 2.4 Secondary antibody controls for immunocytochemical staining: 1 %

BSA/PBS was substituted for primary antibody. (A) Explant derived DPC (B) C6

glioma cells (C) BMSC (D) MC3T3-E1 cells.

A

C

B

D

60

CHAPTER 3 RESULTS

Characterisation and analysis of culture models

3 Characterisation of cell cultures

The primary aim of this study was to investigate the affect of GDNF on DPC

behaviour. For this purpose, different cell models were used; (i) explant-derived and

(ii) enzymatically-derived models of DPC were compared to (iii) BMSC along with (iv)

osteoblast-like MC3T3-E1 cells. The viability of these cell types following exposure to

different culture conditions along with the transcriptional profile and expression of

GDNF receptors, RET and GFRα1 were assessed.

3.1 DPC, BMSC and MC3T3-E1 growth curve analysis under varying culture

conditions

Two in vitro DPC culture approaches were established and compared, these

were explant-derived (Couble et al. 2000) and enzymatically-derived culture (Gronthos

et al. 2000, 2002). DPC were cultured in plastic multiwell plates using 10 %, 20 %

serum, or serum-free conditions at various cell seeding densities and analysed using the

WST-1 assay to determine the number of viable cells. Standard curves obtained

using the WST-1 assay demonstrated a linear relationship between seeding density and

the absorbance of the reduced WST-1 product for both DPC culture models, BMSC and

MC3T3-E1 cells (Figures 3 and 3.1).

61

A

B

Figure 3 Correlation between number of cells seeded and WST-1 formazan

product absorbance. Standard curve for A) explant-derived DPC and B)

enzymatically-derived DPC in serum-free (SF), 10 % FBS and 20 % FBS serum

supplemented conditions. R2 values indicates a linear relationship between seeding

density and WST-1 product absorbance (Mean ± SD; n=3).

R2 =0.9443 R2 =0.9622 R2 =0.9469

R2 =0.8744 R2 =0.8096 R2 =08464

62

Figure 3.1 Correlation between the number of cells seeded and WST-1 formazan

product absorbance. Standard curves for BMSC and MC3T3-E1 cells. Cells were

seeded at a range of densities in 10 % serum supplemented medium. R2 values indicates

a linear relationship between seeding density and WST-1 product absorbance (n=3,

Mean ± SD).

3.1.1 Comparison of growth curves for explant-derived and enzymatically-derived

DPC under serum supplemented conditions after 48 hours

Cells were cultured in 10 % serum supplemented medium for 48 hours to

determine whether changes in cell number remained within a range detectable by the

WST-1 assay at the 48 hour time point. A relationship between seeding density and

absorbance was apparent at 48 hours of culture (Figure 3.1.1). For future experiments an

optimal seeding density of 5000 cells per well (observed at 3.69x just below mid-point

63

on the line graph) was selected to allow decreases and increases in cell number after 48

hours to be within a measureable range for the assay.

Figure 3.1.1 Number of viable explant and enzymatically derived DPC determined

after 48 hours culture in 10% serum supplemented medium. Explant (A) and

enzymatically (B) derived DPC increasing seeding densities increased viable cell

number at 48 hours. Line equations for explant derived DPC and enzmatically derived

DPC were y=-0.2339x2+2.1315x+0.0037 and y=-0.1538x

2+1.6631x-0.0046

respectively. (Explant derived DPC and enzymatically derived DPC number were

determined using the formulae from Figure1; y=0.0141x and y = 0.0263x,

respectively).(n=3, Mean ± SD)( ANOVA = 0.0001 all groups were significantly

different to zero cells seeded).

64

3.1.2 DPC, BMSC and MC3T3-E1 cell viability under serum-free and serum

supplemented conditions

To characterise this culture model further, the viability of explant-derived DPC

were analysed under serum-free culture conditions. Serum is ill-defined, although major

protein constituents are known; batches can differ in various bioactive components,

including growth factors, which contribute to the survival and expansion of cell cultures

(Zolotarjova et al. 2008; Gstraunthaler 2003). Notably, serum supplementation of

media can mask or modify cellular responses during experiments designed to assess the

effects of growth factors (Chase et al. 2010; Wagner & Ho 2007). To establish cell

culture conditions for the culture models under which the effect of GDNF could later be

analysed, the culture of DPC under serum-free conditions was compared to DPC

cultured in 10 % serum supplemented medium at both 24 and 48 hours (Figure 3.1.2).

Based on the reliability of the explant-derived cultures in terms of numbers of cells

obtained per isolation, consistency of cellular morphology, technique reproducibility

and technical speed, the further studies described below utilised this approach.

Serum-free DPC cultures exhibited significantly reduced numbers of viable DPC

compared to 10 % serum cultures (Figure 3.1.2). DPC number did not increase over

time from 24 to 48 hours indicating absence of cell growth under serum-free conditions

(Figure 3.1.2).

65

Figure 3.1.2 Effect of serum-free culture on explant derived DPC at 24 and 48

hours culture. Serum-free (SF) culture for 48 hours significantly decreased the number

of viable explant derived DPC compared to 10 % serum supplemented (10 %) culture

throughout a range of seeding densities (n=3, Mean ± SD;2-way ANOVA * p= 0.003

signifcantly different groups; …

p= 0.0001 significantly different to respective zero cells

seeded).

BMSC and MC3T3-E1 osteoblast-like cells are well characterised models used

for multipotent mesenchymal and osteoblastic cell differentiation, respectively

(Harichandan and Bühring 2011, Jones and McGonagle 2008; Beck et al. 2001; Raouf

and Seth 2000). While the full repertoire and exact integrated regulatory processes

which underpin mineralised tissue formation are not yet completely understood the

transcriptional control and intracellular mechanisms of osteoblast differentiation and

hence bone formation during development and repair are generally better described

within the literature compared with DPC differentiation and dentine formation (Aubin

and Heersche 2000; Karsenty 2008; Robling et al. 2006; Komori 2005; Simon et al.

66

2009; Smith and Lesot 2001; Mitisiadis and Rahiotis 2004; Thelstaf 2006; Lin and

Rosenburg 2011).

BMSC and MC3T3-E1 cultures were therefore used to investigate if differential

responses occur between BMSC, MC3T3-E1 cells and DPC when exposed to the same

culture conditions. The effect of serum-free medium on viable BMSC and MC3T3-E1

numbers at 24 and 48 hours was evaluated (Figure 3.1.3). The number of viable BMSC

and MC3T3-E1 cells did not significantly increase over time following culture in

serum-free conditions. This data was comparable to that obtained for the DPC cultures

(Figure 3.1.2). Indeed a minimal decrease in viable cell DPC and MC3T3-E1 cell

number was apparent after 48 hours under serum-free conditions, although no decrease

in BMSC numbers was evident.

67

A

B

Figure 3.1.3.Effect of serum-free culture conditions on BMSC and MC3T3-E1 cell

growth. Number of viable A) BMSC and B) MC-3T3-E1 (BMSC (n=3), MC3T3-E1

(n=2) Mean ± SD; 2- way ANOVA . * p = 0.001 significant difference from respective

control (zero cells seeded) ; … p = 0.001 significant difference between groups at 24

and 48 hours.

To further investigate whether serum-free conditions would support long-term

differentiation and mineralisation of cultures, the effect of extended culture periods was

68

also analysed. BMSC cultures were not included in this analysis due to the lack of

reduction in viable cell numbers observed at 48 hours compared to 24 hours.

Conversely in serum-free culture at seeding densities of 3125 to 12500 cells per well

BMSC number increased after 48 hours compared to 24 hour culture (Figure 3.1.3A).

Therefore it was on this premise that BMSC could be supported in long term serum-free

culture.

During longer-term cultures (up to 10 days), the number of viable DPC and

MC3T3-E1 cells in serum-free conditions decreased significantly at day 5 (Figure

3.1.4). DPC numbers remained significantly decreased at day 10 compared to day 3.

However at day 10 the decrease in number of viable explant-derived and MC3T3-E1

cells under serum-free conditions was minimal at 16.92 % and 14.43 % respectively.

Figure 3.1.4 Effect of serum-free culture on the number of viable explant-derived

DPC and MC3T3-E1 cells after 3, 5 and 10 days. Viable MC3T3-E1 cell numbers

were signifcantly decreased at Day 5 and viable DPC numbers significantly decreased at

Day 5 and 10. Results are expressed as percentage controls (day 3 value). (n = 6, mean

± SD; TTest …

p= 0.000 and *p= 0.05, resepectively).

*

…

*

69

3.2 Relative gene expression profiles in DPC, BMSC and MC3T3-E1 cultures

3.2.1 GDNF, GFRα1 and RET receptor gene expression within a glial cell line.

To enable the culture models to be screened to detect GDNF and GDNF

receptor (GFRα1 and RET) gene expression, RT-PCR conditions were established using

RNA from rat C6 glioma cell line, which is reported to express both receptors

(Wiesenhofer et al. 2000; Wan and Too 2010). Transcripts for GDNF and its associated

receptors, RET and GFRα1, were detected in accordance with this literature (Figure

3.2).

A B

Figure 3.2 A) sq-RT-PCR analysis of C6 glioma cell line relative gene expression

for the GDNF receptors, RET and GFRα1 and GDNF as analysed by sq-RT-PCR.

GAPDH was used as an amplification control. (B) Representative gel electrophoresis

image of transcripts detected (n=3, Mean ± SD).

70

3.2.2 Relative gene expression of GDNF, Gfrα1 and RET in DPC cultures

Explant-derived and enzymatically-derived DPC culture models were analysed

for the gene expression of GDNF receptors, RET and GFRα1, along with GDNF. The

culture models were additionally analysed for the presence of mesenchymal stem cell

(MSC) marker transcripts, p75NTR

(CD271), CD44 and CD105 (Endoglin) (Pittenger et

al. 1992, 2008; Jones and McGonagle 2008; Bϋhring et al. 2007; Moscatelli et al.

2009) (Figure 3.2.1).

Both DPC models expressed the MSC markers, p75NTR

, CD105 and CD44

(Huang et al. 2009; Strys et al. 2011; Bianco et al. 2001), indicating the mesenchymal

nature of these cultures. GDNF receptor gene expression was evident in both DPC

culture models and in vivo dental pulp and brain tissue. Gene expression levels of

GFRα1 and RET receptor suggested that these receptors may be relatively higher in

explant-derived DPC compared with enzymatically derived DPC (Figure 3.2.1).

71

A B

Figure 3.2.1 sq-RT-PCR analysis of explant-derived and enzymatically-derived

DPC cultures. A) Relative gene expressions of DPC and B) representative

electrophoresis gel images. Brain tissue RNA extracts were used as controls. GAPDH

was used as an amplification control;* p<0.05. (n=4, Mean ± SEM (except CD105 and

CD44 n=2)).

72

These data indicated that the GDNF receptors are expressed in vivo within pulpal tissue

and in vitro in DPC, indicating that both DPC culture models may provide suitable

model systems for the analysis of the effects of GDNF.

3.2.3 Relative gene expression of GDNF, GFRα1 and RET in BMSC and MC3T3-

E1 cell cultures

In vitro cultures of BMSC and the MC3T3-E1 cell line were also investigated

for transcript expression of GDNF and its receptors RET and GFRα1, along with that of

mesenchymal cell markers. Both GDNF and GFRα1 transcripts along with

mesenchymal cell markers were detected within BMSC and MC3T3-E1 cells (Figures

3.2.2 and 3.2.3). Gene expression within BMSC and the MC3T3-E1 cell line indicates

that these models are also potentially responsive to GDNF.

73

A B

Figure 3.2.2 sq-RT-PCR analysis of BMSC relative gene expression for GDNF,

RET & GFRα1 receptors along with mesenchymal cell markers A) Graphical

representation of relative gene expression levels in BMSC and B) representative

electrophoresis gel images of transcripts detected for each gene amplified. GAPDH was

used as an amplification control. (n=4, Mean ±SE).

3.3 Immunocytochemical analysis of GDNF receptor expression

To substantiate the RT-PCR results and determine the protein expression and

localisation of RET and GFRα1, immunocytochemical (ICC) staining was performed

using the C6 glioma cell line as control and in explant-derived DPC, BMSC and

CD105

GFRα1

GDNF

RET

p75NTR

CD44

GAPDH

74

A B

Figure 3.2.3 sq-RT-PCR analysis of MC3T3-E1 gene expression for GDNF, and

RET & GFRα1 receptors along with mesenchymal cell markers. A) Relative gene

expressions of MC3T3-E1 and B) representative electrophoresis gel image. GAPDH

was used as an amplification control (n=3 except for CD44 where n =2; Mean ±SE).

MC3T3-E1 cell cultures (Figures 3.3-3.3.3). Similar uniform distribution of

cytoplasmic staining was visible for the GDNF receptors, RET and GFRα1, within

DPC, BMSC and MC3T3-E1 cell cultures. GFRα1 receptor staining of BMSC and

MC3T3-E1 cells displayed distinct darkly staining areas within the cytoplasm at levels

much elevated above background as seen compared to the control. Specifically, where

staining that appeared to be localised towards the nucleus and may therefore possibly

indicate that the receptors aybe stored within the endoplasmic reticulum within DPC

and MC3T3-E1 cells (Figures 3.3 and 3.3.3).

75

Figure 3.3 C6 Glioma cell cultures stained for the GDNF receptors GFRα1 and

RET cultured in 10 % FBS supplemented medium on multispot glass slides. Non-

specific/negative staining controls used were normal rabbit serum substituted for

primary antibody. The biotinylated secondary antibody was used at 1/200 (column (A))

and higher magnification images are shown in column (B). DAB and Mayer’s

haematoxylin were used for visualisation of antibody binding and as a counterstain,

respectively. Scale bars represent for (A) 100 µm and for (B) 50 µm.

76

Figure 3.3.1 DPC cultures stained for the GDNF receptors GFRα1 and RET in 10

% FBS supplemented medium on multispot glass slides. Non-specific/negative

staining controls used were normal rabbit serum substituted for primary antibody. The

biotinylated secondary antibody was used at 1/200 (column (A)) and higher

magnification images are shown in column (B). DAB and Mayer’s haematoxylin were

used for visualisation of antibody binding and as a counterstain, respectively. Scale

bars represent (A) 100 µm and for (B) 50 µm.

77

Figure 3.3.2 BMSC cultures stained for the GDNF receptors RET and GFRα1 in

10%FBS supplemented medium on multispot glass slides. Non specific/negative

staining controls used were normal rabbit serum substituted for primary antibody. The

biotinylated secondary antibody was used at 1/200 (column (A)) and higher

magnification images are shown in column (B). DAB and Mayer’s haematoxylin were

used for visualisation of antibody binding and as a counterstain, respectively. Scale bars

represent (A) 100 µm and for (B) 50 µm. Arrows depict increased intensity of staining

specific to the receptors within cytoplasm compared to weak diffuse non-specific

controls (normal rabbit serum).

78

Figure 3.3.3 MC3T3-E1 cell cultures stained for the GDNF receptors RET and

GFRα1 in 10% FBS supplemented medium on multispot glass slides. Non-

specific/negative staining controls were normal rabbit serum substituted for primary

antibody. The biotinylated secondary antibody was used at 1/200 (column (A)) and

higher magnification images are shown in column (B). DAB and Mayer’s haematoxylin

were used for visualisation of antibody binding and as a counterstain, respectively.

Scale bars represent (A) 100 µm and for (B) 50 µm. Arrows depict increased intensity

of staining specific to the receptors within cytoplasm compared to weak diffuse non-

specific controls (normal rabbit serum).

79

CHAPTER 4 RESULTS

GDNF effects on DPC, MC3T3-E1 and BMSC survival and proliferation

4 GDNF effects on viable cell numbers

Standard cell cultures contain serum or additional components and growth

factors to support cell viability and growth, whilst serum or growth factor deprivation

often results in apoptosis and senescence. Chapter 3 demonstrated that serum-free

culture decreased DPC and MC3T3-E1 viable cell numbers indicating growth

arrest/increased cell death occurred under these conditions. To study cellular

proliferation, cultures in growth arrest are commonly used to detect a response to

exogenously applied growth factors. In this study, the direct cellular effect of GDNF

was investigated in both serum-free and serum-supplemented cultures.

Changes in viable DPC, BMSC and MC3T3-E1 cell numbers in response to GDNF

were measured under serum-free and serum supplemented conditions using the WST-1

assay. Glioma cell cultures express GNDF receptors and have been reported to be

responsive to GDNF (Ng et al. 2009). Glioma cells were used as a positive control and

GDNF significantly increased the numbers of viable C6 glioma cells under serum-free

culture (Figure 4).

80

Figure 4 GDNF effect on C6 glioma cell numbers as assessed by WST-1 under

serum-free culture conditions after 48 hour culture. Results are expressed as

percentage control (0 ng/ml unsupplemented cultures) (n=3, Mean ± SD; *p< 0.005

versus control).

The addition of GDNF to serum-free DPC and MC3T3-E1 cell cultures also

significantly increased the number of viable cells. Linear regression analysis of

individual experiments for (i) DPC resulted in p < 0.000- 0.007 with R2

values up to

0.49 and (ii) for MC3T3-E1 p>0.000 with R2 values up to 0.54. No effect of GDNF

was apparent in serum supplemented DPC cultures (Figures 4.1). Interestingly, the

number of viable BMSC did not change significantly in response to GDNF addition in

serum-free cultures. However a decrease in viable BMSC and MC3T3-E1 cell numbers

in serum supplemented conditions was evident at 100 ng/ml GDNF (Figure 4.1.1 and

4.1.2).

81

Figure 4.1 GDNF effect on viable DPC number as assessed by WST-1 under

serum-free and 10 % serum supplemented culture conditions. Results are expressed

as percentage controls (0 ng/ml GDNF unsupplemented cultures) (n=4, Mean ± SEM;

*p= 0.0003 versus control).

82

Figure 4.1.1 GDNF effect on viable MC3T3-E1 cell number under serum-free

and 10% serum supplemented culture conditions as assessed by WST-1. Results

are expressed as percentage of controls (n=4, Mean ± SEM; *p= 0.0003 versus control

(0 ng/ml GDNF unsupplemented cultures).

83

Figure 4.1.2 GDNF effect on viable BMSC number under serum-free and serum

supplemented culture conditions as assessed by WST-1 Results are expressed as

percentage of controls (0 ng/ml GDNF unsupplemented cultures) (n=4, Mean±

SEM).

4.1 GDNF effects on cell proliferation and cell viability

To further investigate whether the increase in viable cell numbers was due to increased

proliferation or increased survival, the BrdU assay, and the live/dead assays were

applied.

84

4.1.1 Effect of GDNF on DPC and MC3T3-E1 cell proliferation

A BrdU incorporation assay was performed under serum-free and serum

supplemented conditions for DPC and MC3T3-E1 cells in response to GDNF (100

ng/ml) (Figures 4.2 to 4.2.2). In serum-free media proliferative rates were decreased for

MC3T3-E1 cell cultures compared to cultures containing serum as demonstrated by a

significantly reduced BrdU labelling index (Figures 4.2.2).

At 48 hours, GDNF significantly increased the number of labelled nuclei in

serum-free DPC and MC3T3-E1 cell cultures [BrdU labelled DPC nuclei increased

from 4.5 to 9.9 % and BrdU labelled MC3T3-E1 nuclei from 1.3 to 3 % (Figures 4.2

and 4.2.2]). At 24 hours the numbers of BrdU labelled DPC nuclei under serum-free

conditions with GDNF was similarly increased compared to serum-free conditions

(from 4.5% to 12.2% BrdU labelled DPC nuclei ) (Figures 4.2 ).

Conversely GDNF significantly decreased the number of BrdU labelled DPC

nuclei at 24 hours and MC3T3-E1 BrdU labelled nuclei at 48 hours within serum

supplemented conditions (for DPC, BrdU labelled nuclei decreased from 23.8 % to 5.68

% and for MC3T3-E1 cells, BrdU labelled nuclei decreased from 8.9 % to 5%)

(Figures 4.2.1 and 4.2.2). These data indicate that GDNF increased DPC and MC3T3-

E1 cell proliferation in serum-free cultures whilst decreasing mitosis in serum

supplemented cultures, albeit at differing time points. BMSC cultures were not included

in this investigation because results from the WST-1 assay detected no significant

difference in viable cell numbers following the addition of GDNF in serum or serum-

free cultures at 48 hours (see Figure 4.1.2).

85

A

B

Figure 4.2 BrdU labelling of DPC at 24 and 48 hours under serum-free conditions.

A. Labelled number of DPC nuclei under serum-free conditions with and without

GDNF addition recorded using Image J software at 24 and 48 h. Results are percentage

labelled /total nuclei for each condition (labelling index)(B. Representative microscope

images of stained DPC cultures: BrdU labelled nuclei are shown appearing black and

haematoxylin counterstained nuclei appearing grey. (n=3, Mean ± SEM; *p ≤ 0.05

versus 0 ng/ml GDNF unsupplemented control, scale bars represent 50 µm).

86

A

Figure 4.2.1 BrdU labelling analysis of DPC at 24 and 48 hours under serum

supplemented (10 % FBS) conditions. A. Labelled number of DPC nuclei under

serum supplemented conditions with and without GDNF addition recorded using Image

J software at 24 and 48 h. Results are percentage labelled/total nuclei (labelling index)

for each condition. B. Representative images of stained DPC: BrdU labelled nuclei are

shown appearing black and haematoxylin counterstained nuclei appearing grey. (n=3,

Mean ± SEM; ***p ≤ 0.005 versus 0 ng/ml GDNF unsupplemented control, scale bars

represent 50 µm).

B

87

A

Figure 4.2.2 BrDU labelling analysis of MC3T3-E1 cultures at 48 hours under

serum-free and serum supplemented (10 %FBS) conditions. A. Labelled number of

MC3T3-E1 nuclei under serum-free and serum supplemented conditions with and

without GDNF addition recorded using Image J software at 48 h. Results are

percentage labelled /total labelled nuclei (labelling index) for each condition B.

Representative images of labelled MC3T3-E1 cells: BrdU labelled nuclei are shown

appearing black and haematoxylin counterstained nuclei appearing grey. (n=2, Mean ±

SEM; T-test;*p ≤ 0.05, ***p<0.005 versus 0 ng/ml GDNF unsupplemented control,

scale bars represent 50 µm).

B

88

4.2 Effect of GDNF on DPC, BMSC and MC3T3-E1 cell survival

The live/dead assay was used to assess cell survival in terms of the number of

dead and live cells under serum-free and serum supplemented conditions. A range of

GDNF concentrations were applied under serum-free conditions while 100 ng/ml

GDNF was applied only for serum supplemented conditions. At both 24 and 48 hours

the numbers of dead DPC, BMSC and MC3T3-E1cells under serum-free conditions

without GDNF addition were all significantly increased compared to serum

supplemented cultures (Figures 4.3- 4.3.5). GDNF significantly decreased the

percentage of dead DPC, MC3T3-E1 cells and BMSC under serum-free cultures (albeit

at differing concentrations and time points) (Figures 4.3.1; 4.3.3-4.3.5). DPC death was

significantly decreased at 48 hours at both 10 and 100 ng/ml GDNF supplementation

whereas MC3T3-E1 cell death was significantly decreased at 48 hours in response to 1

ng/ml GDNF (Figures 4.3.1;-4.3.3). GDNF at 10 ng/ml also significantly decreased

BMSC death under serum-free conditions (Figures 4.3.4 and 4.3.5).

Under serum supplemented conditions there were no significant changes in dead

cell numbers with GDNF supplementation, although BMSC consistently demonstrated

reduced levels of cell death at 24 and 48 hours in response to GDNF (Figures 4.3.4 -

4.3.5).

89

A

B

Figure 4.3 Live and dead analysis of DPC under serum-free and serum

supplemented conditions at 24 hours. A) The percentage of dead DPC within each

condition (Serum-free with 0, 1, 10 or 100 ng/ml GDNF and serum supplemented with

0 or 100 ng/ml GDNF). B) Representative images of live and dead staining; arrows

show non-viable cells. Results are expressed as percentage; dead/total DPC. (n=3-7,

Mean ± SEM; …

p = 0.0001 versus 100 ng/ml GDNF under serum-free conditions).

90

A

B

Figure 4.3.1 DPC under serum-free and serum supplemented conditions at 48

hours. A) The percentage of dead DPC within each condition (Serum-free with 0, 1,10

or 100 ng/ml GDNF and serum supplemented with 0 or 100 ng/ml GDNF). B)

Representative images of live and dead staining, arrows show non viable cells. Results

are expressed percentage; dead/total DPC (n=3-7, Mean ± SEM; *p =0.0001 versus

serum supplemented (0 ng/ml GDNF unsupplemented); Δ

p = 0.0001 versus serum-free

control (0 ng/ml GDNF unsupplemented).

91

A

B

Figure 4.3.2 MC3T3-E1 cells under serum-free and serum supplemented

conditions at 24 hours. A) The percentage of dead MC3T3-E1 cells within each

condition (Serum-free with 0, 1,10 or 100 ng/ml GDNF and serum supplemented with

0 or 100 ng/ml GDNF). B) Representative images of live and dead staining, arrows

show non viable cells. Results are expressed as percentage; dead/total MC3T3-E1 (n=3-

7, Mean ± SEM; *p =0.0001 versus serum supplemented (0 ng/ml GDNF

unsupplemented).

92

A

B

Figure 4.3.3 MC3T3-E1 cells under serum-free and serum supplemented

conditions at 48 hours. A) The percentage of dead MC3T3-E1 cells within each

condition (Serum-free with 0, 1, 10 or 100 ng/ml GDNF and serum supplemented with

0 or 100 ng/ml GDNF ). B) Representative images of live and dead staining, arrows

show non viable cells. Results are expressed as percentage; dead/total MC3T3-E1.

(n=3-7, Mean ± SEM; *p =0.0001 versus serum supplemented (0 ng/ml GDNF

unsupplemented); Δ

p = versus serum-free control (0 ng/ml GDNF unsupplemented)).

93

A

B

Figure 4.3.4 BMSC under serum-free and serum supplemented conditions at 24

hours. A) The percentage of dead BMSC within each condition (Serum-free with

0,1,10 or 100ng/ml GDNF and serum supplemented with 0 or 100 ng/ml GDNF). B)

Representative images of live and dead staining, arrows show non viable cells. Results

are percentage; dead/total BMSC. (n=3-7, Mean ± SEM; *p =0.0001 versus serum

supplemented (0 ng/ml GDNF unsupplemented); Δp versus serum-free control (0 ng/ml

GDNF unsupplemented).

94

A

B

Figure 4.3.5 BMSC under serum-free and serum supplemented medium at 48

hours. A) The percentage of dead BMSC within each condition (Serum-free with

0,1,10 or 100 ng/ml GDNF and serum supplemented with 0 or 100 ng/ml GDNF

supplementation). B) Representative images of live and dead staining, arrows show non

viable cells. Results are expressed as percentage; dead/total BMSC. (n=3-7, Mean ±

SEM, *p =0.0001 versus serum supplemented (0 ng/ml GDNF unsupplemented); Δ

p

versus serum-free control (0 ng/ml GDNF unsupplemented).

95

4.2.1 Effect of GDNF on cellular apoptosis and necrosis

To support the live/dead data and detect necrosis and apoptosis within the

cultures further biochemical cell death assays were performed under serum-free

conditions. The LDH (lactate dehydrogenase) cytotoxicity assay determined the overall

levels of cell death from necrosis and/or apoptosis (Xin et al. 2001). At 48 hours

GDNF addition decreased LDH levels for MC3T3-E1 cells compared to (no GDNF)

control or the LDH measurement taken at 24hours. LDH levels for DPC were also

reduced at 48 hours compared to 24 hours. However no significant differences to the

(no GDNF) control cultures were evident for MC3T3-E1 cells, DPC or BMSC (Figure

4.4).

Figure 4.4 Effect of GDNF (100 ng/ml) on LDH levels in supernatants of DPC,

MC3T3-E1 and BMSC cultures at 24 and 48 hours under serum-free conditions.

Control contained no GDNF. Results are expressed as percentage control (0 ng/ml

GDNF unsupplemented (Mean ± SEM, n=3).

96

At 48 hours the activities of caspases-3/-7 as a marker for apoptosis were

investigated within DPC cultures. The activities were significantly increased under

serum-free compared to serum supplemented cultures. Moreover, caspase-3/7 levels

were significantly reduced on GDNF addition compared to control serum-free cultures

containing no GDNF (Figure 4.5). These caspase levels were significantly reduced to

levels comparable to that of serum supplemented cultures (1130 RLU and 1065 RLU,

respectively) indicating that apoptosis was reduced by GDNF supplementation.

Figure 4.5 Caspase 3/7 activities within serum-free DPC cultures. Caspase levels

were significantly reduced after 48 hours culture under serum supplemented conditions

and GDNF (100 ng/ml) supplemented serum-free conditions compared to serum-free

control (0 ng/ml GDNF unsupplemented). Results are shown as relative light units

(RLU) (n=3, Mean ± SEM; T-test *p<0.05).

97

4.3 The effect of attenuating GDNF signalling on DPC and MC3T3-E1 cell viability

GDNF reportedly binds GFRα1, and can then form dimers with the RET

receptors to induce intracellular signalling. (Sariola and Saarma 2003; Kawamoto et al.

2004; Bespalov and Saarma 2007; Airaksinen 1999). To confirm that GDNF action was

direct signalling via its main receptors RET and GFRα1 was reduced using RPI-1 and

PI-PLC inhibitors, respectively (Iwase et al. 2005; Cao et al. 2006; Cuccuru et al.

2004; Lanzi et al. 2000, 2003; Mologni et al. 2005).

4.3.1 Inhibition of GFRα1 (GPI- linked receptor)

PI-PLC hydrolyses GPI-membrane anchors thus cleaving the GFRα1 binding sites from

the membrane. After 48 hours culture with PI-PLC the number of viable cells was

assessed using the WST-1 assay. PI-PLC significantly reduced the number of viable

DPC and MC3T3-E1 cells under serum-free GDNF supplemented conditions at all

concentrations of PI-PLC (from 114 % to 76 % at 0.4 units PI-PLC for DPC and from

144 % to 93 % at 0.2 units PI-PLC for MC3T3-E1 cells), whilst it had no significant

effect at any concentration in control serum-free cultures (Figure 4.6 A and B). The

number of viable GDNF–stimulated DPC and MC3T3-E1 cells was decreased by the

PI-PLC to levels comparable to those of controls at all concentrations of PI-PLC tested.

This data indicated that PI-PLC abrogated GDNF activity on DPC and MC3T3-E1 cells

(Figure 4.16A and B) suggesting that binding to GFRα1 was needed for GDNF effects

on DPC and MC3T3-E1 cells.

98

A

B

Figure 4.6 PI-PLC effect on A) GDNF-stimulated DPC and B) GDNF-stimulated

MC3T3-E1 cells at 48 hours under serum-free conditions. GDNF was added to

cultures at 100 ng/ml. Results are expressed as percentage controls (n=6, Mean ± SD;

*p =0.0001 versus serum-free + GDNF control; …

p=0.01 versus serum-free control (0

ng/ml GDNF unsupplemented).

99

4.3.2 The effect of inhibition of RET receptor DPC and MC3T3-E1 cell viability

RPI-1 competitively inhibits RET receptors by binding at an active site of the

receptor and thereby blocking auto-phosphorylation and subsequent activation of

intracellular signalling pathways (Figure 1.1, pg 4) (Mologni et al. 2005, 2006;

Kawamoto et al. 2004). In serum-free controls, high RPI-1 concentrations (>30 µM)

significantly decreased viable DPC numbers compared to the control cultures indicating

RPI-1 was toxic at these concentrations. Such toxicity has previously been recorded for

this compound (> 30 µM) (Lanzi et al. 2000). At 30-60 µM the RET inhibitor had a

similar cytotoxic effect on MC3T3-E1 cells.

RPI-1 (0.3-3 µM) significantly decreased GDNF-stimulated viable DPC cell

number under serum-free conditions suggesting that GDNF effect was dependent on

signalling through the RET receptor (Figure 4.6.1A). Addition of GDNF with RPI-1 (6

µM) significantly decreased viable MC3T3-E1 cell numbers to levels comparable to the

levels of the serum-free controls (Figure 4.6.1B). The addition of 0.1 ng/ml of GDNF

significantly increased viable glioma cell number (Figure 4). Due to the relatively high

―sensitivity‖ of this cell line to GDNF, the inhibitor was added at higher concentrations

within the range that showed effects on DPC and MC3T3-E1 cells, to ensure adequate

attenuation of the receptors. [ increased sensitivity may indicate increased receptor

numbers (Davidson et al. 2011)]. The addition of the inhibitor at 3 µM and 6 µM

decreased the numbers of glioma viable cells (Figure 4.6.2). This data demonstrated

that RET inhibition reduced the GDNF stimulated increase in the number of viable

100

DPC, MC3T3-E1 and C6 glioma cells under serum-free conditions. These data

indicated that GDNF effects were mediated by the RET receptor for all cell types tested.

A

B

Figure 4.6.1 RPI-1 effect on A) GDNF-stimulated DPC viable cell number and B)

GDNF-stimulated MC3T3-E1 viable cell number at 48 hours under serum-free

conditions. GDNF was added to cultures at 100 ng/ml. Results are expressed as

percentage controls. (n=2, Mean ± SEM, *p =0.0001 versus serum-free + GDNF; ***

p=

versus serum-free control (0 ng/ml GDNF unsupplemented)).

101

Figure 4.6.2 RPI-1 effect on GDNF-stimulated glioma cell number at 48 hours

under serum-free conditions. GDNF was added to cultures at 0.1 ng/ml. Results are

expressed as percentage controls. (n=7, Mean ± SD; *p= 0.0001 versus serum-free

control (0 ng/ml GDNF unsupplemented).

102

CHAPTER 5 RESULTS

GDNF effects on cell differentiation and in vitro mineralisation

5 Short-term in vitro differentiation

The stimulation of DPC, BMSC and MC3T3-E1 cell differentiation during

disease may limit the effects of infection and repair damage by generating cells (i) to

replace lost/damaged cells and (ii) reform the mineralised tissues of the tooth and bone.

In this study alkaline phosphatase (ALP) was used as a marker of early differentiation of

DPC, BMSC and MC3T3-E1 cells in response to GDNF. ALP levels can reportedly be

used as an early indicator of differentiation ( Matsui et al. 1992; Porter et al. 2003;

Beck et al. 1998).

ALP activities were decreased for all cell types studied under serum-free

compared to serum-supplemented culture conditions. Under serum-free conditions,

GDNF (100 ng/ml) significantly decreased BMSC and MC3T3-E1 cell ALP activities

(Figure 5B and 5C). Under serum-supplemented conditions DPC and BMSC displayed

non-significantly reduced ALP activities in response to increasing GDNF concentration.

However changes in MC3T3-E1 cell ALP activities under serum supplemented

conditions were only apparent at 100 ng/ml GDNF where ALP activities were

significantly increased (Figure 5B). This data suggested that GDNF may inhibit early

differentiation of DPC, BMSC and MC3T3-E1cells under serum-free conditions yet

stimulate differentiation of MC3T3-E1 cells under serum supplemented conditions at

100ng/ml (Figure 5).

103

Figure 5 GDNF effect on (A) DPC (B) MC3T3-E1 cells and (C) BMSC ALP

activities under serum-free and serum supplemented conditions. Results are

expressed as percentage of control (0 ng/ml GDNF unsupplemented conditions) (Mean

± SEM; n=4, BMSC and MC3T3-E1 cells, Mean ± SEM; n=3, *p<0.05 versus control).

C

B

A

104

5.1 Long-term in vitro mineralisation

GDNF induced proliferation of DPC and MC3T3-E1 cells at day 2 (see Chapter

4) along with changes in ALP levels shown above (Figure 5). These data suggested that

GDNF may be stimulating early differentiation/expansion within short term cultures

under serum-free conditions and may therefore also affect late differentiation over the

longer term. Notably ECM mineralisation has been reported to be preceded by

transiently increased ALP levels (Hoemann et al. 2009; Wei et al. 2007; Lui et al.

2007).

Staining of mineralised ECM under osteogenic conditions [(supplementation

with 10mM β-glycerophosphate (β-gp), 50 µg/ml ascorbic acid (AA) and 10-7

M

dexamethasone (DEX)] demonstrates that cultures contain fully differentiated

functionally active secretory cells, therefore Alizarin Red Staining (ARS) was used to

visualise and semi-quantify mineralised areas of the cell cultures.

Preliminary mineralisation studies were performed in the absence of GDNF to

determine the capacity of the cell cultures to produce mineralised matrix in standard

serum supplemented cultures. MDPC-23 cells were used as a positive control to

represent odontoblast-like cells known to produce mineralised ECM in response to

osteogenic additives (Hanks et al. 1998; Sun et al. 1998; Pang et al. 2006).

105

Under serum-supplemented osteogenic conditions all cultures showed

significant mineralisation compared to standard serum supplemented conditions that had

minimal mineral deposition (Figure 5.1). MDPC-23 and MC3T3-E1 cells displayed

relatively early mineralised ECM formation and generally stained more intensely

compared to the DPC cultures. Staining of mineralised ECM within MDPC-23 and

MC3T3-E1 cell cultures was evident at day 3, while DPC staining remained

comparatively weak even after 24 days of culture (Figure 5.1). Subsequently the effect

of GDNF was investigated under osteogenic serum-free culture conditions at 2 and 3

week intervals compared to serum supplemented osteogenic conditions to assess GDNF

effects on mineralisation of the cultures.

106

Figure 5.1 Effect of osteogenic culture conditions on in vitro mineralised matrix

formation. Results are shown as percentage of control (day 3), for (A) MDPC-23 cells

(B) DPC and (C) MC3T3-E1 cells. Corresponding images of Alizarin red staining

(ARS) under osteogenic (OSTEO) or control (standard serum supplemented) conditions

are shown on the right. Results are expressed as percentage of day 3 osteogenic

absorbance value (450/630nm; Mean ± SD; n=3, *p <0.005 versus day 3 osteogenic

conditions).

Elu

ted

Aliz

arin

Re

d (

% c

on

tro

l)

Time cultured under osteogenic conditions (Days)

3

107

5.1.1 In vitro mineralisation under serum-free conditions

DPC were cultured under serum-free osteogenic supplemented conditions in

order to study the longer term effects of GDNF. The addition of osteogenic

supplements and GDNF to serum-free conditions significantly increased ARS staining,

compared to serum-free osteogenic cultures and also compared to serum-free cultures

without osteogenic supplements (Figure 5.2). The subsequent mineralisation of the

ECM indicated that GDNF was potentially stimulating osteo/odontogenic

differentiation.

Figure 5.2 Effect of GDNF on mineralised matrix formation for DPC cultures at 7

day. (A) Concentration of eluted Alizarin red (µM) after culture under osteogenic or

serum-free (SF) conditions and (B) corresponding images of Alizarin red staining.

Results are expressed as Alizarin red concentration (µM) (Mean ± SD; n=3, *p<0.001

significantly different groups; p…

versus serum-free unsupplemented controls).

A

B

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The effects of GDNF on in vitro mineralisation were studied using DPC and

MC3T3-E1 cells cultured for 21 and 28 days under serum-free osteogenic conditions

with GDNF, using serum supplemented osteogenic culture conditions as a positive

control for mineralisation. Areas of mineralised ECM were detected within DPC and

MC3T3-E1 cell cultures and the addition of osteogenic supplements to serum-free

conditions increased ARS staining 28 and 21 days of culture respectively. ARS was

significantly increased within GDNF-stimulated cultures under serum-free conditions

(Figure 5.3).

Interestingly when DEX was not used for supplementation the GDNF-stimulated

serum-free DPC osteogenic cultures displayed increased ARS staining indicating that

DEX addition decreased GDNF effects on differentiation. The highest level of staining

was observed for serum supplemented cultures with osteogenic supplements,

furthermore GDNF treated DPC serum-free cultures without DEX showed comparable

staining levels to serum supplemented osteogenic conditions (Figures 5.3).

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A similar pattern of staining was evident for MC3T3-E1 cells, albeit at day 21

osteogenic supplements increased ARS staining under serum-free conditions compared

to controls that contained no osteogenic supplementation. The addition of GDNF to the

osteogenic supplements further increased the intensity of staining detected. In contrast

to the results for DPC, the addition of DEX and GDNF to MC3T3-E1 cell cultures

resulted in staining more intensely than all other conditions at day 21 (Figure 5.4).

These data indicated that DEX and GDNF may act synergistically to increase MC3T3-

E1 cell differentiation. The DPC cultures again stained more weakly under all

conditions compared with MC3T3-E1 cell cultures. At day 28, all osteogenic

supplemented conditions for both DPC and MC3T3-E1 cultures were significantly

increased compared to the controls (Figure 5.4).

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A B

Figure 5.3 Effect of GDNF on in vitro DPC cultures mineralised matrix formation (A) Concentration of eluted Alizarin red (µM)

after culture under osteogenic or control conditions at 21 or 28 days of culture within 48 well plates. Results are expressed as Alizarin

red concentration (µM) (Mean ± SD; n=3, *p<0.05 versus respective control (serum-free or serum supplemented) at time point 21

days or 28 days; ∆p<0.05 significantly different groups at 21 days or 28 days). Results from the 28 day group were not compared to

21 day group (B) corresponding images of Alizarin red staining in triplicate for each condition.

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A B

Figure 5.4 Effect of GDNF on in vitro MC3T3-E1 cell culture mineralised matrix formation. (A) Concentration of eluted Alizarin

red (µM) after culture under osteogenic or control conditions at 21 or 28 days of culture within 48 well plates. Results are expressed as

Alizarin red concentration (µM) (Mean ± SD; n=3, *p<0.000 versus respective control (serum-free or serum supplemented) at time point

21 days or 28 days; ∆p<0.05 significantly different groups at 21 days or 28 days). Results from the 28 day group were not compared to 21

day group. (B) Corresponding images of Alizarin red staining in triplicate for each condition.

* *

*

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CHAPTER 6 RESULTS

Potential role of GDNF during postnatal repair and infection

6 GDNF effects on DPC and/or MC3T3-E1 cell behaviour in response

to;

6.1 Streptococcus mutans (S.mutans)

DPC and MC3T3-E1 cell survival and proliferation may be limited following

exposure to bacteria and in vivo such infections, therefore limit tissue repair and

regeneration. S.mutans is a gram positive bacteria present within the plaque biofilm on

teeth (Michelich et al. 1980; Marsh 2006; Verstraeten et al. 2008) and is associated

with dental caries progression (Rosengren and Winblad 1975; Meikle et al. 1982;

Jackson et al. 1997). S.mutans is therefore regarded as an instigator of tooth attrition

and can contact dental pulp and periodontal bone during aggressive infections.

However, the direct effect of S.mutans on dental pulp cells and bone cells still warrants

further investigation (Hahn and Liewehr 2007; Law et al. 2008; Wayman et al. 1992;

Duany et al. 1971).

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Initially DPC and MC3T3-E1 viable cell number in response to heat inactivated

S. mutans with and without GDNF supplementation were investigated. DPC and

MC3T3-E1 cells were cultured with S. mutans (at 0, 1, 10 and 100 cells per DPC or

MC3T3-E1 cell), with and without 100 ng/ml GDNF and under serum supplemented

conditions. The addition of S. mutans did not significantly affect the numbers of viable

DPC or MC3T3-E1 cells (Figure 6A and B); furthermore GDNF addition had no

significant effect on DPC or MC3T3-E1 cell viability in the presence of heat inactivated

S.mutans (Figure 6A and B).

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A

B

Figure 6 GDNF (100 ng/ml) effect on viable cell number with or without

S. mutans addition for (A) DPC and (B) MC3T3-E1 cell cultures after 48

under serum supplemented conditions. Bacterial cell concentrations were 0,

1, 10 and 100 cells per DPC or MC3T3-E1 cell. Viability was determined

by WST-1 assay. Results are expressed as percentage control (0 ng/ml

GDNF unsupplemented conditions) (Mean ± SD; n=2).

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6.1.1 TNFα

There is potential for the use of the anti-apoptotic properties of GDNF

(Steinkamp et al. 2003; Botchkareva et al. 2000; Dalkara et al. 2011; Ng et al. 2009)

to aid repair of dental pulp and bone in vivo during inflammatory conditions. GDNF

activity under in vitro conditions representative of some aspects of disease may improve

our understanding of GDNF function during dental and bone disease. TNFα is a pro-

inflammatory cytokine, present during infections of the dental pulp and during

inflammatory bone and skeletal joint diseases such as rheumatoid arthritis (Kokkas et

al. 2007; Cooper et al. 2010; Scott and Kingsley 2006). In this study TNFα

supplemented cultures provided a simple in vitro model of aspects of inflammation to

investigate the action of GDNF on cells-derived from mineralised tissue during injury

and inflammation.

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Under serum-free conditions no significant effects of TNFα on viable DPC cell

numbers were detected (Figure 6.1A). The addition of GDNF (100 ng/ml) to the TNFα-

containing cultures increased viable DPC numbers to levels seen for previous

experiments where GDNF alone was added to DPC (Chapter 4). This data indicated that

TNFα displayed no evident interaction with GDNF-induced cell survival and

proliferation under serum-free conditions.

Under serum-supplemented conditions, TNFα (1-100 ng/ml) significantly

decreased viable DPC numbers. Further supplementation with GDNF (100 ng/ml)

abolished TNFα-induced decrease in viable DPC numbers (Figure 6.1B). Conversely,

under both serum-free and serum supplemented conditions, MC3T3-E1 cell numbers

were significantly increased with 10 and 100 ng/ml TNFα supplementation. Moreover

TNFα and GDNF elicited a synergistic effect significantly increasing MC3T3-E1 viable

cell numbers above those obtained from either TNFα or GDNF addition alone (Figure

6.1.1). These data indicate that GDNF may negate the deleterious effects of TNFα on

DPC, whereas for MC3T3-E1 cells GDNF enhances the effects of TNFα.

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A

B

Figure 6.1 TNFα effect on viable DPC number with or without GDNF (100 ng/ml)

supplementation (A) DPC cultured under serum-free conditions and (B) under serum

supplemented conditions for 48hrs. Results are expressed as percentage of controls (0

ng/ml GDNF unsupplemented conditions) (Mean ± SEM; n=3, *p= 0.0001 versus

serum supplemented control; …

p<0.0001 significantly different groups; Δp= 0.032

significantly different groups).

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A

B

Figure 6.1.1 TNFα effect on viable MC3T3-E1 cell number with or

without GDNF (100 ng/ml) supplementation (A) MC3T3-E1 cells

cultured under serum-free conditions and (B) under serum supplemented

conditions for 48h. Results are expressed as percentage controls (0 ng/ml

GDNF unsupplemented conditions) (Mean ± SEM; n=3, *p= 0.0001 versus

serum-free control (*p= 0.023 versus serum supplemented control).

*

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6.1.2 NGF (nerve growth factor)

The neurotrophin, NGF (nerve growth factor) may activate RET, up regulate

GDNF expression and has been detected with GDNF during infection (Dechant et al.

2002; Appel et al. 1997; Martinelli et al. 2006). NGF is also involved in dentine

generation during development and reportedly present during dental pulp injury where it

may promote dentine repair (Mitsiadis and Luukko 1995; Byers et al. 1990; Woodnutt

et al. 2000; Shiomi et al. 2000; O’hara et al. 2009; Arany et al. 2009; Kurihara 2007;

Mizuno 2007). NGF may also be involved in osteoblast survival and bone repair

(Mizumo et al. 2007; Yada et al. 1994; Mogi et al. 2000; Asaumi et al. 2000; Aiga et

al. 2006).

Subsequently, initial experiments were performed to assess the effect of NGF

on DPC viability. NGF significantly reduced viable DPC number under serum-free

conditions at all concentrations analysed and significantly increased viable DPC number

under serum supplemented conditions (Figure 6.2).

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Figure 6.2 NGF effect on viable DPC number under serum-free and 10% serum

supplemented conditions as assessed by WST-1 assay. Results are expressed as

percentage of controls (0 ng/ml NGF, unsupplemented cultures) (Mean± SD; n=6,

*p<0.000 versus control;**p< 0.05 versus control).

In addition, NGF (100 ng/ml) supplementation significantly increased DPC ALP

levels under serum-free conditions; however, no effect was evident under serum

supplemented conditions (Figure 6.2.1). These results indicated that NGF may stimulate

differentiation under serum-free conditions and early differentiation/proliferation under

serum supplemented conditions.

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Figure 6.2.1 NGF effect on ALP activity under serum-free and 10% serum

supplemented conditions. Results are expressed as percentage of controls (0 ng/ml

NGF unsupplemented cultures) (Mean ± SD; n=6, *p<0.007 versus control).

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6.2 Effect of GDNF on short term gene expression in DPC and MC3T3-E1 cells

This study thus far demonstrated that GDNF has effects on viability and

proliferation in both DPC and MC3T3-E1 cell cultures. These effects may be mediated

by several intracellular signalling cascades induced by GDNF-RET-GFRα1 binding

(Airaksinen et al. 1999; Sariola and Saarma 2003). To further investigate the cellular

effects of GDNF, the expression of a panel of genes were analysed where the gene

expression related to: (i) cell proliferation (PCNA, p27kip1), (ii) cell survival/apoptosis

(GDNF, IL-6, p27kip1, TNFα) and (iii) differentiation (Cfbα1, BSP, IL-6). In addition,

the expression of GDNF and associated receptors (GFRα1, RET) were investigated

under the differing culture conditions and in response to GDNF (Figure 6.3. and 6.3.1).

Results from the sqRT-PCR analyses demonstrated clear differences in IL-6

gene expression for DPC following GDNF treatment. In response to GDNF, DPC IL-6

gene expression was increased under serum-free conditions, whereas under serum

supplemented conditions IL-6 expression was significantly decreased. BSP expression

in DPC was also significantly decreased under serum-free conditions compared to

serum supplemented conditions (Figure 6.3). Cbfα1 gene expression was increased for

DPC and MC3T3-E1 cells by GDNF (Figure 6.3 and 6.3.1). In MC3T3-E1 cells BSP

expression exhibited a similar profile to that exhibited by DPC and with BSP decreased

relative levels under serum-free conditions were seen compared to serum supplemented

conditions.

GDNF gene expression was relatively ( albeit non-significantly) increased for

both DPC and MC3T3-E1 cells under serum-free conditions in response to GDNF

stimulation suggesting a possible positive autocrine feedback mechanism was in effect

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(Figure 6.3 and 6.3.1). GFRα1 expression demonstrated the same pattern of expression

as GDNF for both DPC and MC3T3-E1 cells. Unique to DPC cells, GFRα1 appeared

differentially regulated under serum-free compared to serum supplemented conditions

in response to GDNF addition (Figure 6.3). For both DPC and MC3T3-E1 cells RET

receptor expression demonstrated the same profile and was increased under serum-free

conditions compared to serum supplemented conditions. RET gene expression further

decreased in response to GDNF supplementation under both serum-free and serum

supplemented conditions (Figure 6.3 and 6.3.1).

For DPC, TNFα expression appeared increased under serum-free compared with

serum supplemented conditions and GDNF supplementation had a minimal effect on

TNFα expression. p27kip1

expression decreased with GDNF addition in both DPC and

MC3T3-E1 cells under serum supplemented conditions. Under serum-free conditions

GDNF had opposing effects on p27kip1

expression in DPC and MC3T3-E1 cells,

decreasing and increasing in relative levels respectively. PCNA expression

demonstrated a similar pattern of expression in response to GDNF although this was

less evident for MC3T3-E1 cells, where PCNA expression was increased under serum

supplemented conditions compared to serum-free conditions. GDNF addition decreased

PCNA gene expression under serum conditions for both DPC and MC3T3-E1 cells.

Under serum-free conditions relatively small increases in PCNA expression were

detected (Figure 6.3 and 6.3.1).

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Figure 6.3 The effect of GDNF (100 ng/ml) supplementation on relative gene expression in DPC under serum-free and serum

supplemented (10%FBS) conditions at 48 hours. A) Graphical representation of relative gene expression levels and B)

representative electrophoresis gel images of transcripts detected for each gene amplified. GAPDH levels were used as a normalising

control. (Mean ± SEM; n=3, T-test *p<0.05).

A B

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Figure 6.3.1. The effect of GDNF (100 ng/ml) supplementation on relative gene expression of MC3T3-E1 under serum-free and

serum (10 %FBS) supplemented conditions at 48 hours. A) Graphical representation of relative gene expression levels and B)

representative electrophoresis gel images of transcripts detected for each gene amplified. GAPDH levels were used as a

normalising control. (Mean ± SEM; n=2).

A B

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CHAPTER 7 DISCUSSION

7 DPC culture model characterisation

The enzymatically-derived DPC culture model has previously been characterised

in terms of odontoblast differentiation, effect of cryopreservation, pellet culture

techniques, multi-potentiality and ―stemness‖ (Huang et al. 2006; Zhang et al. 2006;

Woods et al. 2009; Yu et al. 2010; Iohara et al. 2004, 2009; Alge et al. 2010; Miura et

al. 2003; Paino et al. 2010). Key papers produced by Gronthos and co-workers

furthered interest in the regenerative capacity of the dental pulp by identifying putative

stem cells within it. Gronthos et al. (2000, 2001, 2002, 2006) found that enzymatically-

derived DPCs could be expanded for up to 20-30 population doublings and were

capable of differentiating along various cell lineages as well as forming a dentine-pulp

complex in vivo when these cells were transplanted into immune-compromised mice. In

these and more recent studies, DPC have been isolated by digestion using trypsin for up

to 30 mins (Patel et al. 2009). This approach, effective for deriving many different cell

types from the ECM, released cells from the dental pulp by cleavage of cell surface

peptide bonds on the C-side of Arginine and Lysine residues (Gilbert et al. 2006).

Enzymatic digestion may therefore significantly disrupt cellular membranes and affect

viability, reducing cell yields (Kirkpatrick 1985). Indeed yield per isolation is a critical

factor when potentially only a proportion of clonogenic DPC (possibly < 20 %) are

capable of more than 20 doublings (Gronthos et al. 2002; Mao et al. 2006).

Couble et al. (2000) applied an explant approach to obtain DPC, identifying

differentiated odontoblast-like cells at four weeks within confluent cultures following

addition of ascorbic acid and β-glycerophosphate to the media. This method has since

127

been used by several groups to study the interaction of these cells with bacteria, dental

materials, pulpal innervation and ECM (Staquet et al. 2008; Farges et al. 2011; Durand

et al. 2006; Maurin et al. 2004; Seux et al. 1991; About et al. 2000; Carrouel et al.

2008).

The use of bone explants to obtain human osteoblast cultures emerged in the late

1970’s and these osteoblast-like cells have since been extensively characterised (Jones

and Boyde 1977; Ecarot-Charrier et al.. 1983; Gallagher et al. 2003; Lian and Stein

1992, 2003, 2004, 2006). Other techniques for isolating cells from mineralised tissues

have also been developed (Iohara et al. 2004; Sasaki et al. 2008; Tjäderhane et al.

1998; Téclès et al. 2008). Cao et al. (2006) found a short period of enzymatic digestion

followed by the explanting technique produced cells more quickly compared with the

enzymatic culturing technique when used for isolating osteoblasts. Technically the

explant approach comprises fewer steps, is less time consuming and requires minimal

manipulation of the tissue compared to the enzymatic digestion method and in the

study presented here this approach was found to produce consistent results. Although

enzymatic digestion approaches may result in cell damage or derivation of less robust

cells, a more heterogeneous cell population containing DPC at different levels of

differentiation may be obtained which may be more representative of the native pulpal

tissue (Patel et al.. 2009; Balic and Mina 2010; Balic et al. 2010).

In this study, explant outgrowth resulted in cells of homogenous polygonal

morphology although this approach took several hours longer to obtain cells compared

with the enzymatically released DPC approach (Figure 2.1). The explant approach

proved to be more efficient, where the reuse of explants was possible thereby producing

further DPC outgrowths (Couble et al. 2000). Therefore the explant-derived model,

128

albeit a more ―selective‖ approach, also appears an appropriate and suitable method for

obtaining DPC.

7.1 GDNF and GDNF receptor expression in DPC cultures

To further characterise the explant and enzymatically-derived cells used in this

study relative gene expression levels were compared. Data presented here demonstrate

that explant-derived DPC displayed a different profile of GDNF, GFRα1 and RET

mRNA expression compared with enzyme-digested DPC, however both were suitable

models to study GDNF affects on DPC in vitro. Indeed explant-derived DPC expressed

relatively higher expression levels of both RET and GFRα1 receptors compared with

enzymatically derived DPC. This differential gene expression suggested that different

cell types may have been selected according to the isolation procedure. However further

qPCR is needed for precise quantification of transcript levels involved. Indeed

differential gene and protein expression according to the isolation procedure used has

previously been reported for neuronal, smooth muscle and dental pulp cell cultures

(Iohara et al. 2004; Arthur et al. 2008; Zimmerman 2006; Kirschenlohr et al. 1995;

Huang et al. 2006b). Zimmermann et al. (2006) found that osteopontin levels were

significantly increased within explanted compared to enzymatically digested smooth

muscle cell cultures. In addition DPC gene expression level for collagen type I was

increased while DSP levels decreased, in enzyme-digested compared with explant-

derived DPC cultures (Huang et al. 2006). Furthermore Zhang et al. (2005) found 5 %

of the DPC population expressed the STRO-1 antigen (a putative mesenchymal stem

cell marker) within explanted rat cultures whereas Laino et al. (2006) found only 9.98

% STRO-1 antigen positive cells within enzymatically isolated DPC.

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Interestingly while RET is reported to have extremely low postnatal expression,

in vitro expression was detectable within DPC cultures. RET expression can represent

the degree of differentiation of some cell types, for example during neuronal

differentiation RET expression increases as cells become more differentiated (Ibanez

and Ernfors 2007; Hoffman et al. 2005; Hoffman 2008). It is conceivable that the

relative difference in gene expression of GDNF and the GDNF receptors between

explanted and enzymatically-derived cell cultures may represent the presence of cell

populations exhibiting varying degrees of differentiation.

7.1.1 Confirmation of the mesenchymal nature of DPC culture models

RT-PCR analysis indicated that the dental pulp and DPC cultures consisted of

cells of mesenchymal nature as evidenced by the expression of the putative MSC

markers p75NTR

(CD271), CD44 (hyaluronan receptor) and CD105 (endoglin is also a

TGF-β co-receptor) (Pittenger et al. 1992, 2008; Jones et al. 2002; Bϋhring et al.

2007; MoscatelliI et al. 2009). p75NTR

expression in DPC was relatively low compared

with dental pulp tissue and this may have reflected the lack of neuronal cells present

within these cultures. p75NTR

is regarded as a putative stem cell marker (Kendirci et al.

2010; Stemple and Anderson 1992; Moscatelli et al. 2009) and its expression may also

indicate the presence of multipotent cells within cultures (Gronthos et al. 2000).

Notably the MSC markers (CD44, CD105 and p75NTR) have been used to isolate

multipotent MSC from pulpal tissue using FACS and these enriched sub-populations

can subsequently form mineralised ECM in vitro (Huang et al. .2009, Bianco et al.

2001).

130

GFL (Glial Family Ligands) are known survival factors for neural crest and

neural crest-derived cells (Enomoto et al. 2005; Sariola and Saarma 2003; Chalazonitis

et al. 1998 Bunone et al. 2000; Insua et al. 2003; Hofmann et al. 2005). The dental

papilla and hence DPC are of neural crest origin and the expression of GDNF, RET and

GFRα1 may reflect this neural crest lineage (Saavedraa 2008; Paratcha and Ledda

2008). GDNF gene expression as detected in this study concurs with the demonstration

of GDNF in rat tooth by Western blot analysis (Kvinnsland et al. 2004) and GDNF

mRNA localisation within the inner dental epithelium (IDE), enamel knots, sub-

odontoblast and odontoblast layers (Nosrat et al. 1998; 2002). GFRα1 and RET were

also expressed in the epithelial and mesenchymal dental papilla, respectively, during

tooth morphogenesis (Luukko et al. 1997; Nosrat et al. 1997) suggesting a role in the

reciprocal epithelial-mesenchymal interactions which occur during early tooth

development. However this study now demonstrates for the first time the GDNF

receptor expression occurring within in vitro DPC cultures.

7.1.2 Characterisation of bone cell cultures for GDNF and GDNF receptor

expression

The co-expression of RET and GFRα1 in BMSC and osteoblast-like cells as

shown here is also a novel observation. GDNF mRNA and protein expression have

previously been detected in BMSC cultures (Ye et al. 2005; Chen et al. 2007; Garcia et

al. 2004) and this study confirmed the expression of GDNF within MC3T3-E1

osteoblast-like cells suggesting a role for GDNF in osteoblast regulation. GDNF

transcript expression in BMSC was shown to be relatively weak, although

immunocytochemical staining was clearly apparent. Differences between the level of

gene and surface protein expression may occur and could be due to post-transcriptional

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and post-translational modifications/processing. Indeed GDNF receptor protein

expression is influenced by post-translational modification, as 30% of translated RET

may misfold in the endoplasmic reticulum and subsequently be degraded (Abrescia et

al. 2005, Kjaer and Ibàñez 2003). Further work based on these studies is now

warranted to analyse the in situ distribution of GDNF and GDNF receptors in whole

tooth and bone tissue. It should also be noted that the expression of the GDNF receptors

Gfrα1 and RET does not necessarily denote responsiveness to GDNF. Indeed

intracellular cross-talk with other growth factor receptor systems occurs whereby TGF-

β is required for RET receptor activation (Tsui-Pierchala et al. 2002; Dechant 2002;

Airaksien 1999; Sariola and Saarma 2003). Nonetheless, the data from the studies

presented here demonstrated that GDNF exerted direct effects mediated by Gfrα1 and

RET on non-neuronal cell types from the dental pulp, calvarial bone and to a lesser

extent the bone marrow (see below).

7.1.3 In vitro culture under serum-free conditions

Serum-free cultures facilitate the study of the direct effects on cells without

interference from serum factors, but also present a model mimicking pathological

conditions involving cellular insult and injury due to growth factor deprivation

(Goyeneche et al. 2006). In this study DPC and MC3T3-E1 cell cultures displayed

reduced cell viability when cultured under serum-free conditions in agreement with

other reports (Onisha et al. 1999; Tarle et al. 2010; Tumber et al. 2000). Interestingly,

BMSC cultures retained relatively good viability which corresponded with previous

studies showing that serum-free conditions did not affect viable BMSC numbers during

a 7 day experimental period (Sauerzweig et al. 2009; Pochampally et al. 2004).

Interestingly Sauerzweig et al. (2009) identified those BMSC proliferating under

132

standard serum-free conditions as differentiating towards a primitive neural lineage. It

would therefore be interesting to investigate whether BMSC can respond to subsequent

GDNF addition by differentiating along a neural cell lineage.

Serum deprivation is known to arrest mitosis and serum starved cultures have

been shown to exhibit differential gene expressions compared with serum supplemented

cell cultures (Raff et al. 1992; Ye and Lotan 2008). Interestingly Onisha et al. (1999)

have previously reported that canine DPC could not be cultured under serum-free

conditions without additional supplements. Reportedly cell-cell and cell-ECM

interactions are likely important in regulating survival under stressful conditions. Indeed

fibronectin (FN) coating of culture surfaces decreased the number of floating, apoptotic

SHED (Stem cells from human exfoliated deciduous teeth) under serum-free conditions

(Tarle et al. 2010) indicating that the loss of cell attachment under these conditions

could be rescued by cell-ECM contact. In addition, differentiated osteoblasts secrete FN

(Stephansson 2002; Garcia and Reyes 2005; Globus et al. 1998; Moursi et al. 1996)

and apoptosis was induced in MC3T3-E1 cells by neutralising FN binding (Globus et

al. 1998). BMSC also synthesise FN constitutively in vitro and FN is important in

BMSC cell-cell adhesion in vivo; however, additional FN coating of BMSC culture

surfaces had no effect on BMSC adherence (Jo et al. 2011; Lerat et al. 1993; Van der

Velde-Zimmermann et al. 1997). Combined this may suggest that variations in

endogenously secreted ECM such as FN may be responsible for differences in cell

viability of DPC and BMSCs under serum-free culture conditions.

In future, investigations utilising sub-optimal conditions for BMSC culture may

be useful to study the positive effects of mitogens or survival factors such as GDNF on

BMSC. Sub-optimal conditions may be established by reducing seeding density or

133

perhaps blocking FN adherence using antibodies against integrin β1 binding sites in

serum-free culture conditions. Data presented here indicated that although DPC and

MC3T3-E1 cell viability was reduced at day 1 and 2 sufficient cell numbers remained

after 10 days of culture, indicating that serum-free conditions can be used to support

long-term studies.

7.2. Short term effect of GDNF on cell proliferation under serum-free conditions

This study demonstrated that GDNF increased DPC and MC3T3-E1 viable cell

numbers and cell proliferation under serum-free conditions. The increase in DPC viable

cell numbers by GDNF corresponded with a decrease in p27kip1

expression confirming

that cell replication/cycle progression was evident and GDNF regulation of p27kip1

expression was in line with other reports (Kalechman et al. 2003; Baldassarre et al.

2002). GDNF did not however affect p27kip1

expression in the osteoblast-like cells

suggesting that this may represent a cell type-specific effect. PCNA (proliferating cell

nuclear factor) is also a marker and regulator of cell cycle progression and cell

proliferation (Maga and Hubscher et al. 2003; Coltrera and Gown 1991; Sasaki et al.

1992; Golberg et al. 2008). In this study PCNA transcript expression was detected as

being both up- and down-regulated in accordance with increasing and decreasing

proliferation as assessed by BrdU incorporation, thus confirming the GDNF-induced

changes in proliferation under serum-free conditions.

7.2.1 GDNF and cell survival

GDNF is a well characterised cell survival factor (Enomoto et al. 2005; Sariola

and Saarma 2003; Chalazonitis et al. 1998 Bunone et al. 2000; Insua et al. 2003;

Hofmann et al. 2005). It is known that the cellular repertoire involved in cell-cycle

134

control and hence proliferation is closely associated with pro-survival signalling

(Maddika et al. 2007). To assess whether the increased viable cell numbers evident in

response to GDNF were also due to increased survival of DPC and MC3T3-E1 cells,

live/dead, LDH and caspase assays were performed to assess the levels of cell death and

apoptosis within all cultures. GDNF decreased numbers of dead DPC that were

observed under serum-free conditions. Moreover, caspase levels indicated that apoptosis

occurred within serum-free DPC cultures and that this was reduced in response to

GDNF addition. This finding is in agreement with other studies demonstrating that

serum withdrawal induces cell death in neuronal cell cultures and this is abrogated by

GDNF supplementation (Kobori et al. 2006). Further work is needed to confirm and

visualise apoptosis, e.g. using the TUNEL assay. In addition, the anti-apoptotic effect

elicited by GDNF remains to be confirmed for the MC3T3-E1 cell cultures.

Interestingly, up-regulation of NTGF receptors under serum-free conditions in the

absence of ligand resulted in apoptosis in several cell types (Ahn et al. 2005; Fauchais

et al. 2008; Yuan and Yanker 2000). Within this study RET and GFRα1 expression

were detected under all culture conditions tested and the inhibitor studies indicated that

the functional effects were mediated by GDNF receptor binding.

7.2.2 Osteogenic differentiation and in vitro mineralisation

Cbfα1/runx2 gene expression in DPC and MC3T3-E1 cells was up-regulated by

GDNF under serum-free conditions; moreover relative expression levels of Cbfα1 were

increased in DPCs under serum-free compared with serum-supplemented conditions.

Cbfα1 is considered a pivotal osteoblast transcription factor, although it may not be

essential for odontoblast differentiation (Narayanan et al. 2001; Fen et al. 2002; Qin et

al. 2007). Cbfα1 is used as a marker of the osteoblast lineage and may reflect the

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osteogenic potentiality of cells in culture (Galindo et al. 2005; Wang et al. 1999).

Moreover, ALP activities in the DPC, BMSC and MC3T3-E1 cell cultures were

decreased by GDNF supplementation under serum-free conditions, suggesting an

inverse relationship between cell proliferation and differentiation (Yokose et al. 2000;

Hoemann et al. 2009; Maddika et al. 2007).

An in vitro mineralisation assay was conducted as a measure of functional

osteogenic differentiation. Cultures were grown for up to 28 days under in vitro osteo-

inductive conditions and stained with alizarin red (ARS) to detect mineralised ECM

under serum-free conditions. Initial experiments indicated that mineralisation occurred

at a later time point for DPC compared to MC3T3-E1 cells suggesting that DPC

cultures were less differentiated or less inductive under these study conditions compared

with the osteoblast-like cell line. The relatively long-term mineralisation experiments

indicated that GDNF stimulated osteogenic differentiation of both DPC and MC3T3-E1

cell cultures. It may be possible that cell death occurring under serum-free conditions

related to the formation of mineralised ECM. Lynch et al. (1998) found apoptosis was

co-regulated with osteoblast differentiation and Fratzl-Zelman et al. (1998) found rapid

mineralisation occurred within cultures containing dead cells. However, other studies

have reported mineralisation within osteoblast and BMSC cultures under serum-free

conditions (Gronowicz et al. 1989; Sakamoto et al. 1989; Huffman et al. 2007;

Solmesky et al. 2010). In this study the mineralisation observed under serum-free

conditions with osteogenic induction became significant only after 28 days; thus it is

improbable that this calcification was associated with dead cells which normally occurs

relatively rapidly within the early stages of culture (Fratzl-Zelman et al. 1998).

136

The level of ARS under serum-free osteogenic conditions was changed with the

addition of GDNF and/or removal of DEX. GDNF without DEX supplementation in

MC3T3-E1 cell cultures decreased ARS staining, whereas ARS staining was increased

in DPC cultures, indicating cell specific effects of DEX in combination with GDNF.

The mechanism of action for GDNF increased ECM calcification remains unclear.

GDNF may stimulate early osteogenic/odontogenic differentiation of mesenchymal

cells and/or expansion of existing progenitor cells; moreover, GDNF may exert late

effects on the synthesis of ECM proteins (Del Rio et al. 2011; Lin al. 2001). GDNF

may also have promoted endogenous production of growth factors such as BMPs that

subsequently enhanced cell differentiation during the in vitro mineralisation

experiments. Indeed GDNF may act synergistically with BMP-4 and -7 during neuronal

development and proliferation (Chalazonitis et al. 2004; Harvey et al. 2005).

7.3 Short-term effect of GDNF on cell proliferation and differentiation under

serum-supplemented conditions

GDNF decreased viable MC3T3-E1 cell number by inhibiting proliferation

under serum supplemented conditions as confirmed using a BrdU assay. This study also

demonstrated that relative gene expression of the osteogenic-associated transcription

factor Cbfα1 was up-regulated in MC3T3-E1 cell cultures under serum supplemented

conditions by GDNF. GDNF is known to induce differentiation not only for neuronal

cell types, but also for cells of the testes and kidney (Peterson et al. 2004; Toshifumi et

al. 2005; Baldassarre et al. 2002; Meng et al. 2000; Insua et al. 2003; Park et al.

2005). In this study relative gene expression of BSP in the osteoblast cultures was

enhanced under serum supplemented conditions. Furthermore the decrease in

proliferation in GDNF supplemented cultures may also be indicative of

137

cytodifferentiation (Aubin and Heersche 2000; Blagosklonny 2003). Interestingly,

GDNF/RET signaling has been shown to be responsible for decreased proliferation of

an embryonic neural precursor carcinoma cell line (Baldassarre et al. 2002). This

effect, mediated by p27kip1

, was suggested to be a mechanism by which GDNF regulates

cell growth to initiate terminal differentiation (Baldassarre et al. 2002). Moreover the

short-term experiments performed in this study demonstrated that ALP levels were

increased concomitantly with decreasing proliferation. Further long-term studies will be

required to investigate in detail the role of GDNF in osteogenic proliferation,

differentiation and bone formation.

7.4 In vitro culture under “challenging” conditions that reflect some aspects of

infection and inflammation

In vivo bacterial infection or injury to bone and dentine can lead to the

proteolysis of the ECM and elicit an inflammatory reaction by the host immune system.

Enhanced levels of GDNF found in gingival crevicular fluid from patients with chronic

periodontitis emphasize the potential involvement of GDNF in the pathophysiology of

dental tissues (Sakai et al. 2006). Previous studies have demonstrated that LPS

(lipopolysaccharide) and NFĸB (a TNFα regulated transcription factor) signalling up-

regulated NGF expression during dental inflammation (Freund-Michel and Fossard

2008; Heese et al. 1998; Magloire et al. 2001). The cytokine TNFα is well known for

its ability to stimulate recruitment and activation of immune cells to the sites of

infection (Zganiacz et al. 2004; Abe et al. 2010; Stashenko et al. 1991; Stashenko et

al. 1987; Li and Stashenko 1992), however it is also reportedly able to stimulate

osteogenic migration, proliferation and differentiation (Gowen et al. 1988; Frost et al.

1997; Glass et al. 2011; Lencel et al. 2011; Modrowski et al. 1995). TNFα also directs

138

the differentiation of osteoclasts that resorb bone, although the exact mechanisms by

which this occurs are not yet fully ascertained (Robinson et al. 2007; Nanes 2003).

Given GDNF effects on osteoblast-like and odontoblast-like cell differentiation and

survival under serum-starved conditions, the effect of GDNF under challenged

conditions recapitulating some aspects of infection and inflammation were investigated.

7.4.1 Streptococcus mutans (S.mutans) exposure

S.mutans is a gram positive bacterium associated with dental caries and

Rosengren and Winblad (1975) demonstrated that S.mutans infections are also involved

in deeper destruction of bone during dental infections. However, the effects of S.mutans

on the viability of the dental pulp or osteoblasts are not elucidated. Meikle et al. (1982)

found S.mutans antigens contributed to inhibition of MC3T3-E1 cell protein synthesis

and that this may contribute to bone loss. In accordance, bacterial wall components of

S.mutans and LTAs (Lipoteichoic acid) have been found to induce bone resorption

(Nair et al. 1996).

Elson et al. (2007) studied the direct effect of heat inactivated gram positive and

negative bacteria compared with purified LPS on TLR activation and concluded that

heat inactivation was a valid method for studying TLR responses of the innate immune

system. Previously, Paterson et al. (1982, 1987) have indicated that live S.mutans

decreased dental pulp cell numbers in vivo; however decreasing cell viability was

evident from day 7 only suggesting that this was not a direct cytotoxic effect of the

bacteria. More recently Abe et al. (2010) reported heat inactivated S.mutans stimulated

DPC osteogenic and dentinogenic differentiation. Interestingly the literature suggests

that GFL expression during bacterial infection may increase cell survival, whereby

139

infecting gram positive bacteria express outer protein structures that mimic NTGF

structure and subsequent exposure to the host cells has been reported to increase

neuronal cell differentiation and survival during bacterial infection (Lu and

PereiraPerrin 2008). Moreover, Durand et al. (2006) found that LTA exposed

odontoblasts up-regulated mRNA for hypoxia-inducible factor 1α (HIF1α), a

transcription factor regulating GDNF expression (Foti et al. 2010). Within this study no

effect of heat inactivated S.mutans on viable DPC or MC3T3-E1 cell number was

evident. Likewise S.mutans had no evident effect on GDNF stimulated DPC and

MC3T3-E1 viable cell number. Further work is needed to assess S.mutans effect in

isolation from serum factors and to determine whether bacterial components modulate

GDNF effects under these conditions.

7.4.2 GDNF interaction with TNFα

Interestingly the NFĸB transcription factor has a binding site on the human

GDNF promoter, suggesting a potential regulatory feedback loop involving GDNF in

response to inflammatory signalling (Appel et al. 1997; Woodbury et al. 1998). Under

serum-free conditions TNFα exerted no effect on viable DPC numbers and GDNF

addition increased viable cell numbers to levels previously seen without TNFα addition.

Thus in vitro experiments were conducted using serum to mimic the multitude of

proteins present under physiological conditions. This study demonstrated that under

serum supplemented conditions, TNFα decreased DPC viable cell numbers, and this

effect was abated by GDNF addition. These data suggested that GDNF may protect

DPC against TNFα induced apoptosis/necrosis in agreement with previous reports

(Talley et al. 1995; Yang et al. 2002; Sheng et al. 2005; Peralta-Solter et al. 1996).

140

Interestingly, TNFα promoted osteoblast proliferation which was further

enhanced in the presence of GDNF. The mechanisms by which GDNF interacted with

TNFα to stimulate osteoblast proliferation are as yet unknown, but plausibly the effect

may have been due to the stimulation of converging pathways that may include CREB

(cAMP response element), IKK and NFĸB signalling (Ono et al. 2004; Kalechman et

al. 2003; Encinas et al. 2008). Furthermore TNFα also is proposed to regulate GDNF

expression and secretion. (Tanaka et al. 2000; Woodbury et al. 1998; Kim et al. 2009;

Simon et al. 2007; Esseghir et al. 2007; Kuno et al.. 2006; Appel et al. 1997). Thus, it

is possible that increased GDNF synthesis by TNFα could lead to autocrine positive

feedback mechanisms.

7.4.3 Cell specific effects of GDNF

This study demonstrated that whilst GDNF had direct effects on DPC and

MC3T3-E1 cell viability, GDNF only decreased ALP levels in BMSC cultures under

serum supplemented conditions but had no obvious comparable effect on BMSC

proliferation or viability. These data corresponds with previous findings by Kramer et

al. (2006). These apparent cell-specific effects of GDNF may reflect cell-specific

receptor expression and intracellular signalling (Encinas et al. 2008; Tsui-Pierchala et

al. 2002; Encinas et al. 2001; Focke et al. 2001; Mason et al. 2006) and/or may be

due to the differential expression of tissue specific genes such as PAX and SOX genes

in long bones as opposed to neural crest derived tissues (Fernandas et al. 2004; Betters

et al. 2010; Dezawa et al. 2005; Otto et al . 2009; Basch et al. 2006; Wang et al .

2009). Indeed DPC and MC3T3-E1 cells displayed similar proliferative responses to

GDNF. Furthermore it should be noted that MC3T3-E1 cells are of foetal origin

whereas BMSC are postnatally derived cells. Gattei et al. (1997) hypothesised that

141

GDNF has a role within the bone marrow, activating signalling between BMSC

expressing GFRα1 and RET expressing hematopoietic cells to stimulate monocytes

differentiation.

This study highlighted another novel action of GDNF, that the neurotrophic

factor modified cytokine IL-6 gene expression in DPC and MC3T3-E1 cell cultures.

These findings are consistent with previous studies indicating a role for IL-6 in late

stage differentiation and survival of osteoblasts and fracture healing (Jüttler et al. 2002;

Li et al. 2010; Jilka et al. 1998; Iwasaki et al . 2008; Tsiridis and Giannoudis 2006). IL-

6 is also a known neurokine able to sustain neuronal cell survival (Hirano et al. 2000;

Bromberg and Wang 2009) and it may therefore be speculated that IL-6 may play a role

downstream of GDNF as described in the current study.

7.4.4 NGF stimulation of DPC

It was previously proposed that the neurotrophin, NGF, may be involved in

odontoblast differentiation and dentine formation during tertiary dentinogenesis (Arany

et al. 2009; Kurihara 2007; Woodnutt et al. 2000; Shiomi et al. 2000; O’hara et al.

2009). Neurotrophins have been detected during fracture repair and NGF may also be

involved in the repair of bone (Asaumi et al. 2000; Aiga et al. 2006). NGF was reported

to stimulate differentiation and survival of osteoblasts (Mizumo et al. 2007; Yada et al.

1994; Mogi et al. 2000). NGF can also up-regulate GDNF expression (Appel et al.1997)

and may modulate its action in vivo during infection and inflammation (Martinelli et al.

2006). This study demonstrated NGF increased DPC viable cell numbers under serum

supplemented conditions. Conversely, NGF decreased viable DPC numbers and

increased ALP activity under serum-free conditions suggesting NGF stimulated cell

142

differentiation. Further studies are required to investigate the inter-relationship between

GDNF and NGF in the regulation of DPC and osteoblast activity.

7.5. Summary

In summary, this PhD study demonstrated that BMSC and MC3T3-E1 cells

which are well characterised models for osteoblast differentiation expressed both GDNF

receptors (GFRα1 and RET). Dental pulp and dental pulp-derived cell cultures also

express the GDNF receptors. Furthermore, direct cell-specific, receptor-mediated

effects of GDNF were demonstrated on the proliferation and cell survival of DPC

cultures. However, GDNF did not affect BMSC proliferation or viability nor did GDNF

influence MC3T3-E1 cell survival under serum-starved conditions. GDNF effects on

DPC and osteoblast cell proliferation were inversely related to ALP levels. Nonetheless,

GDNF stimulated in vitro mineralisation in long-term DPC and MC3T3-E1 cultures.

Further work is warranted to investigate the precise effects of GDNF on osteoblast and

odontoblast differentiation, the mechanisms involved in GDNF signalling in these

mesenchymal/NCC-derived cells and whether GDNF may be used as a bioactive

therapeutic agent in bone or dental repair.

143

CHAPTER 8 CONCLUSION

In conclusion the findings from this research demonstrate that GDNF is able to

directly stimulate odontoblast and osteoblast-like cell proliferation and differentiation.

From these findings, it is proposed that GDNF may play an important role in the

regulation of dental pulp homeostasis and bone metabolism. Moreover GDNF promoted

DPC survival under ―challenged conditions‖ indicating that GDNF may be involved in

the survival and differentiation of DPC and possibly odontoblasts during the repair of

mineralised tissues.

8.1 Clinical and therapeutic applications

The putative role for GDNF during development and homeostasis, regulating

cell differentiation, survival and proliferation may be exploited for clinical regenerative

or therapeutic applications.

8.1.2. GDNF as a bioactive in tissue repair and regeneration

GDNF has already been used within clinical trials for neuronal diseases and

GDNF regulation may be involved in rare disease such as Waardenburg-Hirschsprung

diseases (Pelet et al. 1998; Angrist et al. 1996; Chan et al. 2003). GDNF receptor

signalling may be also involved in neuro-endocrine cancers, breast cancers, leukaemias

and drug dependence (Morandi et al. 2011; Plaza-Menacho et al. 2010; Lanzi et al.

2003, 2009; Gattei et al. 1999; Niwa et al. 2007; Petrangolini et al. 2006). Indeed RPI-

1 and other indoline blockers that inhibit RET have been proposed as clinical treatments

for such diseases (Morandi et al. 2011; Phay and Shah 2010).

144

Non-steroidal anti-inflammatory drugs (NSAIDS) are commonly used to treat

bone diseases such as osteoarthritis, rheumatoid arthritis, ankylosing spondylitis and

osteomyelitis, however their effects are limited (Donnelly and Hawkley 1997; Harder

and An 2003). Anti-TNFα therapy is a promising new treatment however this has many

reported serious side effects (Bongartz et al. 2006; Williams et al. 1992; Vaz et al.

2009). GDNF appears to modify TNFα effects and may represent a target for inhibition

in cases where anti-TNFα treatment is used. However for this application GDNF

regulation and signalling requires further elucidation. Moreover GDNF interactions

with TNFα, highlight a GDNF/TNFα synergy in stimulating osteoblast differentiation

which could possibly be harnessed to promote bone formation. MSC have previously

been genetically engineered and used to deliver growth factors as a part of gene therapy

and similar investigations regarding the dental pulp have been proposed to stimulate

natural reparative events in situ (Moutsatsos et al. 2001; Nakashima et al. 2002,

Kimelman et al. 2007). BMSC and DPSC have also gained much interest for use in

neuronal regeneration due to DPC release of NTGF and DPC have been implanted in

vivo to deliver NTGF or differentiate along neuronal lineages (Bolliet et al. 2008;

Huang et al. 2009; Nosrat et al. 2001; Lillesaar et al. 2001, 2003; Apel et al. 2009).

There is now scope within the field of dentistry for the addition of bioactives such as

GDNF. Bioactives may be used within scaffolds, dental sealants/fillings and via

implantation of bioactive secreting MSC to encourage regeneration of the pulp by

increasing adherence (to biomaterials such as implants/scaffolds), survival,

proliferation, differentiation and migration of dental pulp cells along with promoting

angiogenesis (Brydone et al. 2010; Hing 2004; Zuk et al. 2008; Sun et al. 2010). This

145

study raises the possibility that GDNF may play a role in MSC survival, recruitment and

proliferation during dentine repair (Shi et al. 2008).

8.1.3 Role of GDNF in tooth inflammation and analgesia

GDNF may protect against TNFα induced cytotoxicity during painful pulpal

inflammation where high levels of TNFα occur such as during irreversible pulpitis

(Kokkas et al. 2007; Donaldson 2006). Current treatment for pain relief during

irreversible pulpitis involves tooth extraction or pulp removal (pulpectomy/extirpation)

(Komabayashi and Zhu 2010). GDNF has recently been reported to have analgesic

effects on pain induced by damaged nerves (Takasu et al. 2011; Adler et al. 2009).

Therefore novel dental pulp clinical therapies using GDNF can be envisaged which

maintain and promote tooth vitality (Komabayashi and Zhu 2010; Wang et al. 2010,

2011).

146

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APPENDIX

PUBLISHED WORK

Gale Z, Cooper P.R and Scheven B.A.A (2011) Effects of GDNF on dental pulp cells.

Journal of Dental Research. 90(10):1240-5

Gale Z, Cooper PR, Scheven BA (2012) Glial cell line-derived neurotrophic factor

influences proliferation of osteoblastic cells. Cytokine. 57(2):276-81

ABSTRACTS/CONFERENCE PAPER

Gale Z, Cooper P.R and Scheven B.A.A Glial Cell Line-Derived Neurotrophic Factor

Influences Osteoblast Proliferation. British Society of Oral and Dental Research

Conference Sheffield University. 12-15th

September 2011

Gale Z, Cooper P.R and Scheven B.A.A. Survival of dental pulp stromal/stem cells in

response to Glial Cell Line-Derived Neurotrophic Factor (GDNF) Festival of Research

and Enterprise, University of Birmingham 4-5th

April 2011

Gale Z, Cooper P.R and Scheven B.A.A. Effects of GDNF on dental pulp cells.

International Association for Dental Research, Barcelona July 14-17th 2010

Gale Z, Cooper P.R and Scheven B.A.A. Regulation of dental pulp cells by GDNF.

College of Medical and Dental Sciences Research and Enterprise Gala 19-20th April.

University of Birmingham 2010

Gale Z, Cooper P.R and Scheven B.A.A. Festival of Research and Enterprise Poster

Conference, University of Birmingham. December 2008

PRIZES

Highly commended poster prize (2011) College of Medical and Dental Sciences

Research and Enterprise Gala. University of Birmingham. 4-5th

April

British Society of Oral and Dental Research MINTIG prize (Mineralised Tissue

Group) (2010) IADR. Barcelona. July 14-17th

Medici Prize for best poster in the category enterprise and innovation (2008) Festival of

Research and Enterprise. University of Birmingham. December

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AUTHORS PERSONAL COPIES OF PUBLISHED WORK

PAPER 1

Glial Cell Line-Derived Neurotrophic Factor Promotes Survival and

Proliferation of Dental Pulp Cells

Zoe Gale, Paul R. Cooper, and Ben A.A Scheven

School of Dentistry,

College of Medical and Dental Sciences

University of Birmingham,

Birmingham, U.K.

ABSTRACT

This study investigated the effects of glial cell line-derived neurotrophic factor (GDNF)

on dental pulp stromal cells (DPCs). Cultures of DPCs expressed GDNF as well as its

receptors, GFR1 and RET. Addition of recombinant GDNF to cultures in serum-

containing medium did not significantly affect DPC growth; however, GNDF dose-

dependently increased viable cell number under serum-free culture conditions.

Live/dead, lactate dehydrogenase (LDH) and caspase -3,-7 assays demonstrated that cell

death occurred under serum-free conditions, and that GDNF significantly reduced the

number of dead cells by inhibiting apoptotic cell death. GDNF also stimulated cell

proliferation in serum-free conditions, as assessed by the BrdU incorporation assay. The

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effect of GDNF was abolished in the presence of specific inhibitors to GFR1 and RET

suggesting receptor-mediated events. This study also demonstrated that GDNF

counteracted TNF-induced DPC cytotoxicity, suggesting that GDNF may be

cytoptotective under disease conditions. In conclusion, our findings indicate that GDNF

promotes cell survival and proliferation of DPC and suggest that GDNF may play a

multi-functional role in the regulation of dental pulp homeostasis.

Key words: pulp biology, neurotrophic factors, GDNF, TNF alpha, cell survival, cell

proliferation.

INTRODUCTION

Neurotrophic factors have been implicated in the development and regulation of dental

tissues ( Luukko et al., 1997; Nosrat et al., 1998, 2002; Woodnutt et al., Magloire et al.,

2001). The glial cell line-derived neurotrophic factor (GDNF) is part of GDNF family

of ligands (GFLs) which include neurturin (NRTN), artemin (ARTN) and persephin

(PSPN). GFLs are considered to belong to the TGFβ superfamily sharing partial amino-

acid sequence homology and structural confirmation (see for review Airaksinen and

Saarma, 2002). GDNF is a soluble signaling molecule that binds to a specific membrane

receptor, the GDNF family receptor (GFR1) which forms complexes with the

tyrosine kinase receptor RET or alternative co-receptors such as NICAM eliciting

intracellular signals for cell growth and differentiation (Sariola and Saarma, 2003).

GDNF was originally characterized as a potent trophic factor promoting the survival

and differentiation of neurons (Airaksinen and Saarma, 2002). GDNF was subsequently

shown to be expressed in various tissues outside the nervous system and an important

functional role has been recognized in urogenital tissues, in particular relating to kidney

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development and spermatogenesis (Sariola and Saarma, 2003). The expression of

neurotrophic factors within the adult dental pulp highlighted that GDNF may be

involved in neuronal innervations, axon growth and function (Nosrat et al., 1997; Fried

et al., 2000; Lillesaar et al., 2001; Luukko et al., 1997)). During tooth development,

GDNF and its receptors GFR1 and RET are transiently localized in dental organ

epithelium and pulpal mesenchyme suggesting a role for GDNF in epithelial-

mesenchymal interactions (Hellmich et al., 1996; Luukko et al., 1997; Nosrat et al.,

1998)). Interestingly, ultrastructural analysis of molar tooth germs from GDNF-

knockout mice revealed that ameloblast and odontoblast differentiation was disrupted

suggesting a role for GDNF during tooth cytodifferentiation ((de Vicente et al., 2002)).

We postulated that GDNF may be involved in the regulation and maintenance of the

postnatal dentin-pulp complex. The aim of this study was to investigate the direct

effects of GDNF on dental pulp cells (DPC).

MATERIALS & METHODS

Dental pulp cell (DPC) cultures

DPC cultures were established from rat dental pulp explants as described previously (

e.g., Couble et al., 2000; Huang et al., 2006). In brief, dental pulp was extracted from

incisors of 4-6 week-old male Wistar rats and dissected into small (~5mm3 ) samples

and cultured in tissue culture flasks containing αMEM supplemented with 20% Fetal

Bovine Serum (FBS), 1% penicillin/streptomycin and 2.5 μg/ ml Amphotericin B

(Sigma Aldrich, UK) in a humidified 5% CO2 incubator at 37 OC. DPC proliferated

from the explants, showing a polygonal stromal/fibroblast-like morphology which

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became more cuboidal when reaching confluency. Subconfluent cell cultures were

trypsinized with Trypsin/EDTA (Invitrogen/Gibco, UK) and subcultured. DPC at

passage 2 to 4 were used for the experiments. DPC were seeded into 96-multiwell

plates in MEM/10% FBS at 5,000 cells/well. After 24 hours, the cultures were

replenished with either serum (10% FBS) or serum-free MEM supplemented with

0.1% bovine serum albumin (BSA). Recombinant human GDNF (rhGDNF, Amgen,

Thousand Oaks, USA) was added to the cultures for a further two days. Additional

experiments included recombinant human tumour necrosis factor-α (TNFα; Peprotech,

UK) which was co-incubated with 100 ng/ml GDNF in αMEM/10%FBS for 2 days

before analysis. For the receptor inhibitor experiments, DPC cultures were treated for 1

hour with different concentrations of phosphoinositide phopsholipase C (PI-PLC;

Sigma) which blocks signalling via GFRα1 (Krieglstein et al., 1998) , or RPI-1

(Merck/Calbiochem), a specific RET receptor tyrosine kinase inhibitor (Cuccuru et al.,

2004) followed by further culture with the respective inhibitors in media with or without

GDNF.

Cell number and viability assays

We used the WST1 assay (Roche Applied Biosciences) to assess the number of viable

cells (Scheven et al., 2009); the absorbance of the reduced compound was measured at a

wavelength of 450 with a reference filter at 630nm using a Biotek plate reader. To

distinguish between the number of viable and dead cells in the cultures, we performed a

―live/dead‖ assay using acridine orange (4µM) (stains nuclei of live cells) and ethidium

bromide (4µM) (labels nuclei of dead cells). The numbers of live and dead cells per

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microscopic field was counted under a Nikon Eclipse fluorescent microscope using

480nm and 520nm filters, respectively.

Cell death and apoptosis assays

The level of cell death in the cultures was determined biochemically using a lactate

dehydrogenase (LDH) cytotoxicity assay (Roche, UK). Cell culture supernatants were

analysed after a 2-day culture for the presence of LDH. Absorbance was determined at

490/630nm using the Biotek plate reader. To determine the level of cellular apoptosis,

we used the Caspase-Glo-3/-7 to determine the enzymatic activities of caspsase-3/-7

(Promega); the cleaved luminescent product was measured with a Berthold microplate

luminometer.

BrdU cell proliferation assay

Cell proliferation was assessed with the use of a 5-Bromo-2-deoxy-uridine (BrdU)

labelling and detection kit (Roche Applied Sciences). In brief, cells were labelled with

10 µM BrdU for the final hour of the 48-hour culture followed by fixation and

immunostaining for BrdU incorporation using a specific anti-BrdU antibody. Cells were

counterstained with hematoxylin and the total number of labelled and non-labelled

nuclei were counted in 50 independent microscope fields.

Semi-quantitative RT-PCR (sqRT-PCR) analysis: DPC cultures as well as dental

pulp and brain extracted from 4-week-old rats underwent lysis in RLT buffer

containing β-mercaptoethanol followed by RNA isolation using the RNeasy minikit

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(Qiagen, UK). Subsequently, 1-2 μg of DNase-digested total RNA was used for

oligo(dT) (Ambion, UK) reverse transcription to generate single-stranded cDNA using

the Omniscript kit (Qiagen,UK). Centrifugal filters (Microcon) were used to purify and

concentrate resultant cDNA. Both RNA and cDNA concentrations were determined

from absorbance values at a wavelength of 260 nm using a BioPhotometer (Eppendorf,

UK). sqRT-PCR assays were performed using the RedTaq PCR system (Sigma, UK)

and the Mastercycler gradient thermal cycler (Eppendorf, UK). Primers were designed

from NCBI mRNA sequences using Primer-3 design software (Table 1).

Immunocytochemistry

DPC were seeded onto multispot microscope slides and incubated for 24h at 37 0C in a

humidified 5% CO2 incubator (20,000 cells/well). Rat glioma C6 cells known to express

GDNF as well as GFRα1 and RET (Song and Moon, 2006) were used as positive

controls. The adherent cells were fixed with ice-cold acetone for 5 min, then rinsed in

phosphate-buffered saline (PBS) containing 1%BSA. Following incubation in 3% H2O2

for 30 min (to block endogenous peroxidase) the slides were washed in PBS/1%BSA

and incubated in 20% normal goat serum followed by incubation with 2 µg/ml primary

polygonal rabbit antibody against GFRα1 (sc10716, SantaCruz) or against RET (sc167;

SantaCruz) overnight at 4 0C. The GFRα1 antibody (sc10716, Santa Cruz, USA)

specifically binds to the 58-kDa protein, as determined by immunoblotting, while the

RET antibody (sc167; Santa Cruz) recognizes the 150- to 179-kDa protein (Pierchala et

al., 2006); both antibodies have been validated for use in immunocytochemical staining

of cell membrane receptors (Alladi et al., 2010; see also manufacturer’s datasheets at

http://www.scbt.com/datasheet-10716-gfralpha-1-h-70-antibody.html;

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http://www.scbt.com/datasheet-167-ret-c-19-antibody.html). For controls, the primary

antibody was substituted normal rabbit serum. The slides were rinsed in PBS/1%BSA

and labelled and stained with biotin-streptavidin-HRP using a Biogenex detection kit

(LP000-UL). The slides were counterstained with haematoxylin before examination

using a Zeiss microscope.

Data and statistical analysis

Data obtained from the WST-1, LDH and caspase-3/7 assays were corrected for

background values and expressed as percentage of controls. Data were analysed by

ANOVA with Tukey post hoc test.

RESULTS

DPC culture model

RT-PCR analysis showed that GDNF and its receptors GFRα1 and RET transcripts

were present in postnatal dental pulp and DPC cultures (Fig. 1A). Immunocytochemical

staining of DPC using specific antibodies against GFRα1 and RET suggested the

presence of these GDNF receptors (Fig. 1B). These data indicated that the cultures

provided a suitable model for study of the direct effects of GDNF on DPC.

GDNF stimulates DPSC survival and proliferation

We performed initial experiments to determine the effects of GDNF on cell growth

using serum-containing or serum-free cultures. GDNF did not elicit significant changes

in the number of viable DPCs over a 2-day culture in medium supplemented with FBS;

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however, addition of GDNF to serum-free cultures resulted in a dose-dependent

increase in viable DPC numbers (Fig. 2A). We performed experiments to evaluate

whether the GDNF-induced increase in cell numbers was due to increased cell survival

and/or cell proliferation.

The live-dead assay demonstrated that significant cell death occurred in the control,

serum-free DPC cultures as compared to serum-supplemented cultures (Fig 2B). The

presence of GDNF significantly increased total cell number, coinciding with a

significant decrease in the number of dead cells in serum-free cultures, indicating that

GDNF promoted cell survival under these conditions. (Fig. 2B).

To further characterise the effect of GDNF on cell survival, we performed a

biochemical cytotoxicity assay to measure LDH release from damaged and dying cells.

The results show that GDNF significantly reduced LDH secretion in these cultures (Fig.

2C). To determine if the protective effects of GDNF were due to prevention of cellular

apoptosis, we determined cellular levels of caspase-3/7. Results demonstrate that

caspases-3/7 levels were significantly reduced in GDNF-treated cultures compared with

control cultures, indicating GDNF prevented DPC apoptosis (Fig. 2C).

Next, cell proliferation was determined using by the BrdU-incorporation assay. The

results showed that GDNF significantly increased the number of BrdU-labeled DPC in

serum-free cultures, demonstrating that, along with a pro-survival action, GDNF

stimulated cell replication (Fig. 2D).

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Receptor-mediated effects of GDNF

To determine whether the GDNF effects described above were mediated via its

canonical receptors, we treated cultures with PI-PLC, which cleaves the receptor

subunits from their glycosylphosphatidylinositol (GPI)-anchored membrane proteins.

Analysis of the data presented in Fig. 3 demonstrated that PI-PLC abolished GDNF

effects on viable cell numbers in serum-free cultures, suggesting an essential role for

GFRα1 in the GDNF effects on DPC viability (Fig. 3A). Moreover, RPI-1, a

competitive ATP-dependent RET kinase inhibitor, dose dependently blocked GDNF

action, underscoring GDNF dependency on the RET co-receptor (Fig. 3B).

GDNF counteracts TNFα-induced reduction in DPSC number

Finally, this study investigated whether GDNF may have cell- protective effects under

conditions that better reflected the pathological environment. For this purpose, DPCs

were cultured in serum-containing medium (to obtain an optimal physiological milieu)

which was supplemented with TNFα, a pro-inflammatory cytotoxic cytokine up-

regulated in pulpitis (McLachlan et al., 2004). The results demonstrated that TNFα

dose-dependently decreased DPC numbers; however, cultures supplemented with

GDNF showed significantly increased DPC viability as compared with TNFα controls,

indicating that GDNF counteracted TNFα-induced cytotoxicity (Fig. 4).

DISCUSSION

The notion that neurotrophic factors are implicated in tooth development and dentin-

pulp biology is not surprising due to the cranial neural crest cell origin of dental pulp

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mesenchymal stem cells and odontoblasts and the close association of these cells with

the pulp neuronal network (Fried et al., 2000; Tziafas et al., 2000). Dental pulp is

increasingly gaining attention as a therapeutic tool in nerve repair and regeneration, due

to its neurogenic potential and endogenous expression of neurotrophic factors (Nosrat et

al., 2001; Lillesaar et al., 2001; Apel et al., 2009). The neurotrophic factor GDNF was

shown to be expressed in both ecto-mesenchymal dental papilla as well as inner dental

epithelial cells. GDNF is therefore implicated as an important factor controlling

epithelial-mesenchymal interactions during tooth development, and in

GDNF-knockout mice, ameloblasts and odontoblasts fail to differentiate fully (Hellmich

et al., 1996; de Vicente et al., 2002). GDNF and its canonical receptors GFRα1 and

RET are also expressed in dental pulp and (sub) odontoblasts in postnatal teeth,

suggesting a role in odontoblast function (Nosrat et al., 1997, 1998; Luukko et al.,

1997). This study reports the expression of GDNF and its specific receptors, GFRα1

and RET in dental pulp and DPC cultures and provides evidence that GDNF is a pro-

survival growth factor for DPC via interaction with the GFRα1/RET receptor complex,

suggesting that GDNF may have a functional role in the regulation of dental pulp cells.

Our results corroborate the well established role of GDNF as a cell survival factor and

regulatory signaling factor for neuronal and non-neuronal cells (Airaksinen and Saarma,

2002; Sariola and Saarma 2003). Serum-free cultures facilitate the study of direct

effects on cells without interference of serum factors, but also present a model

mimicking pathological conditions involving cellular insult and injury due to trophic

factor deprivation (Goyeneche et al., 2006). Serum withdrawal induces toxicity in

neuronal cell cultures which is abated by GDNF (Kobori et al.,2006).

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In this study, GDNF was shown to inhibit DPC death induced by serum deprivation,

suggesting that GDNF may play a cytoprotective role in dental pulp homeostasis during

stress conditions and pulpal necrosis. Furthermore, the current paper offers the first

evidence that GDNF is able to block cytotoxic effects on DPC by the pro-inflammatory

cytokine TNFα, highlighting a possible protective role of this neurotrophic factor in

pulpitis. TNFα is able to elicit various cellular responses, depending on cell types and

conditions; in particular, TNFα’s cytotoxic effects are widely documented as involving

apoptosis-related pathways mediated by its main TNF receptor TNFR1 (Shen and

Pervaiz, 2006). The cytotoxic effects of TNFα found in our DPC cultures may involve

induction of cell death combined with an inhibitory effect on cell proliferation. This

finding corresponds with a previous study describing the cytoprotective effects of

GDNF in an adrenal cell line undergoing TRAIL (TNFα-related apoptosis-inducing

ligand)-induced cell death (Murata et al., 2006). Interestingly, TNFα induced production

of GDNF by astrocytes and glioma cells possibly via the NFkβ binding present on the

human GDNF gene promoter, suggesting a regulatory ―protective‖ feedback loop

involving GDNF in response to inflammation (Appel et al., 1997; Woodbury et al.,

1998). The enhanced levels of GDNF found in gingival crevicular fluid from patients

with chronic periodontitis emphasize the potential involvement of GDNF in the

pathophysiology of dental tissues (Sakai et al., 2006).

Apart from its effect on cell survival, GDNF was shown here to be a mitogen for DPCs.

This is not a very surprising finding as it is well known that the cellular repertoire

involved in pro-survival signaling is closely associated with cell cycle control processes

(Maddika et al., 2007). Our results raise the possibility that GDNF may a role in MSC

211

recruitment and proliferation during dentine repair (Shi et al., 2008). GDNF is able to

bind extracellular proteoglycan heparin sulphate chains (Rider, 2006), indicating the

likelihood that GDNF produced by odontoblasts is sequestered within dentin (Nosrat et

al., 1997, 1998). Indeed, preliminary findings from our laboratory using an antibody

array technology have demonstrated that GDNF was detected within human dentine

matrix extracts (Thompson et al., unpublished observations).

This suggests that odontoblast-secreted GDNF can be sequestered within the dentin

could be released from the matrix upon injury or disease. It is noteworthy that some of

the neurotrophic effects of GDNF require the presence of TGF which induces the

translocation of GFR1 to the plasma membrane (Krieglstein et al., 1998; Peterziel et

al., 2002). However, a limitation of the current study is that it is not possible to

conclude which particular dental pulp cell type(s) responded to GDNF. The DPC

cultures cannot be considered homogenous, although, due to the nature of the culture

method (explant-outgrowth of cells), the DPCs generally displayed a morphologically

similar polygonal fibroblast/stromal-like cell appearance. Previously, DPCs have been

shown to express several mesenchymal stem cell features and the capability to

differentiate along different mesenchymal lineages, including osteogenic and

odontogenic lines (see also Couble et al., 2000; Huang et al., 2006). Considering that

GDNF and its receptors are expressed in postnatal dental pulp as well as

(sub)odontoblasts (Luukko et al., 1997; Nosrat et al., 1997; this study), it is plausible to

speculate that GDNF acts upon cells of mesenchymal origin, including those of the

odontoblast lineage, and that GDNF in concert with other local signaling factors such as

TGFβ1 may control cell viability and recruitment during reparative processes within the

dentin-pulp (Tziafas et al., 2000; Woodnutt et al., 2000; Magloire et al., 2001).

212

In conclusion, this study demonstrates that GDNF promoted cell survival and

proliferation of DPC under serum-starved or pro-inflammatory conditions. We propose

that GDNF may have mutli-functionality within the dentin-pulp complex, acting as both

survival factor and mitogen during tooth injury and repair. Further studies are warranted

to evaluate the role of GDNF in dental pulp homeostasis and its potential in novel

therapeutic strategies for dental pulp repair and tissue regeneration.

ACKNOWLEDGEMENTS

This study was supported by a University of Birmingham School of Dentistry PhD

research grant and was awarded the MINTIG prize at the 2010 IADR meeting in

Barcelona. We are grateful to Amgen Ltd (Thousands Oaks, USA) for the supply of the

recombinant GDNF.

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215

Tables and Figures

Gene Primer sequence Accession no. Product Tm

GAPDH F-CCCATCACCATCTTCCAGGAGC

R-CCAGTGAGCTTCCCGTTCAGC NM017008 450bp 60

GDNF F-AGAGGAATCGGCAGGCTGCAGCTG

R-AGATACATCCACATCGTTTAGCGG NM019139 337bp 60

RET

F-TCAGGCATTTTGCAGCTATG

R-TGCAAAGGATGTGAAAGCAG NM001110099 393bp 62.5

GFRα1 F-AATGCAATTCAAGCCTTTGG

R-TGTGTGCTACCCGACACATT U59486 218bp 60

Table 1. PCR primer sequences and annealing temperatures (Tm)

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Figure 1: A. RT-PCR gel images showing transcripts of GDNF and its receptors

GFRα1 and RET in dental pulp mesenchymal cells (DPCs), dental pulp and brain. B.

Immunocytochemical staining of DPSC for GFRα1 and RET. Non-immune rabbit

serum without specific primary antibody was used as control.

217

Figure 2. A. Effect of GDNF on viable cell number in serum-supplemented or serum-

free cultures as assessed by WST-1. Results are expressed as percentage of controls

(mean + SD; n=4). B. Effect of GDNF on the number of dead cells as determined by the

live/dead assay. Results show percentage of dead cells (mean + SD; n=3) C. Relative

LDH and caspase-3/-7 levels after 2-days culture in serum-free medium cultures

supplemented with 100 ng/ml GDNF (mean + SD; n=3). GDNF-treated DPC cultures

showed significantly reduced LDH and caspase-3/7 levels indicating GDNF-induced

cell survival effects involves at least in part an anti-apoptotic effect by GDNF D. BrdU

incorporation after 2-day DPC culture in serum-free medium. The proportion of cells

labeled positively for BrdU (labeling index) was significantly increased in GDNF-

supplemented cultures indicating stimulation of replication by GDNF. Statistical

differences versus serum-free controls: * p < 0.05, **P<0.01, *** p<0.001.

218

Figure 3: Effects of GDNF receptor inhibitors on GDNF-stimulated DPC cultures. A.

Effect of phosphatidylinositol-specific phospholipase C (PI-PLC) on viable cell number

as assessed by WST-1 in cultures supplemented with 100 ng/ml GDNF. B. Effects of

the RET kinase inhibitor, RPI-1, on viable cell number in 2-day GDNF-treated DPC

cultures. Results are percentage of control values as determined by the WST1 assay

(mean + SD; n=6-8). * p < 0.05, **P<0.01 versus GDNF cultures.

219

Figure 4: Effects of 100 ng/ml GDNF on 2-day DPC cultures in the presence of

increasing concentrations of TNFα. Results are percentage of control values determined

by the WST1 assay (mean + SD; n=5-6). *P< 0.05, **P<0.01 versus corresponding

TNFα cultures.

220

PAPER 2

Glial cell line-derived neurotrophic factor induces proliferation of

osteoblastic cells

Zoe Gale, Paul R. Cooper, and Ben A. Scheven

School of Dentistry, College of Medical and Dental Sciences,

University of Birmingham, Birmingham, U.K.

ABSTRACT

Little is known about the role of neurotrophic growth factors in bone metabolism. This

study investigated the short-term effects of glial cell line-derived neurotrophic factor

(GDNF) on calvarial-derived MC3T3-E1 osteoblasts. MC3T3-E1 expressed GDNF as

well as its canonical receptors, GFR1 and RET. Addition of recombinant GDNF to

cultures in serum-containing medium modestly inhibited cell growth at high

concentrations; however, under serum-free culture conditions GDNF dose-dependently

increased cell proliferation. GDNF effects on cell growth were inversely correlated with

its effect on alkaline phosphatase (ALP) activity showing a significant dose-dependent

inhibition of relative ALP activity with increasing concentrations of GDNF in serum-

free culture medium. Live/dead and lactate dehydrogenase assays demonstrated GDNF

did not significantly affect cell death or survival under serum-containing and serum-free

conditions. The effect of GDNF on cell growth was abolished in the presence of

inhibitors to GFR1 and RET indicating that GDNF stimulated calvarial osteoblasts via

221

its canonical receptors. Finally, this study found that GDNF synergistically increased

tumor necrosis factor-α (TNF-α)-stimulated MC3T3-E1 cell growth suggesting that

GDNF interacted with TNF-α-induced signaling in osteoblastic cells. In conclusion, this

study provides evidence for a direct, receptor-mediated effect of GDNF on osteoblasts

highlighting a novel role for GDNF in bone physiology.

Key words: GDNF, neurotrophic factor, TNF-alpha, osteoblast, bone, calvarial, cell

proliferation

1. Introduction

Glial cell line-derived neurotrophic factor (GDNF) is a pleiotropic signaling molecule

playing a pivotal role in the development and regulation of the nervous system (1-2).

GDNF has been recognized as a potent survival factor for neuronal cells in addition to

its essential roles in neural migration and differentiation (1-3). GDNF is also widely

expressed outside neuronal tissues and has been suggested to be involved in epithelial-

mesenchymal interactions during development of urogenital and dental tissues (1, 4-6).

GDNF is able to elicit various intracellular signalling cascades via multiple receptor

systems, primarily through the glycosyl-phosphatidylinositol-anchored, GDNF family

receptor (GFR1) and the tyrosine kinase transmembrane co-receptor RET (2-3).

Neurotrophic growth factors and cytokines including GDNF have been shown to be

expressed in bone marrow stromal cells prompting an emerging interest in therapeutic

regenerative application of bone marrow-derived mesenchymal stem cells in

neurological disorders (7-11). Interestingly, whilst GFR1 expression was detected

222

along with GDNF in bone marrow stromal cells, RET proved to be absent in these cells

(12-13). However, GDNF/GFR1 complexes cleaved from the stromal cells were

shown to elicit functional signaling through RET expressed on hematopoietic and

leukemic cells suggesting a signaling pathway involving cell-cell interactions within the

bone marrow environment (13-14). In this study we addressed the question whether

GDNF may be involved in bone metabolism. In particular, this study focused on the

short term effects of GDNF on the proliferation and survival of osteoblastic cells using

a non-transformed calvarial-derived cell line as model system for osteoblasts. In

addition, the research investigated a possible interaction between GDNF and the

multifunctional, pro-inflammatory cytokine, tumor necrosis factor-α (TNF-α).

2. Materials and methods

2.1 Cell cultures

The MC3T3-E1 cell line is a non-transformed, clonal osteoblast-like cell line

established from mouse calvaria and has extensively been used as a physiologically

relevant in vitro model for calvarial osteoblasts, osteogenic differentiation and bone

formation (15). MC3T3-E1 were acquired from the European Collection of Cell

Cultures (ECACC) and were cultured in αMEM containing 10% Fetal Bovine Serum

(FBS), 1% penicillin/streptomycin, 200 mM glutamine and 2.5 μg/ ml Amphotericin B

(Sigma Aldrich, UK) in a humidified 5% CO2 incubator at 37o

C. Subconfluent cell

cultures were trypsinised using Trypsin/EDTA (Gibco, UK) and plated into 96-

multiwell plates in MEM/10% FBS at 5,000 cells/well. After 24 hours, the cultures

223

were replenished with either serum (10% FBS) or serum-free MEM supplemented

with 0.1% bovine serum albumin (BSA). Recombinant human GDNF (rhGDNF,

provided by Amgen, Thousand Oaks, USA) or TNF-α (PeproTech, UK) was added to

the cultures for a further two days. For the receptor inhibitor experiments, cultures were

treated for 1 hour with different concentrations of phosphoinositide phopsholipase C

(PI-PLC; Sigma) which blocks signalling via GFRα1 (16), or RPI-1

(Merck/Calbiochem), a specific RET receptor tyrosine kinase inhibitor (17), followed

by further culture with the respective inhibitors in media with or without GDNF.

2.2 Cell number and viability assays

The WST1 assay (Roche Applied Biosciences) was used to assess the number of viable

cells (18); the absorbance of the reduced compound was measured at a wavelength of

450 with a reference filter at 630nm using a Biotek plate reader. The ―live/dead‖ assay

used 4µM acridine orange to stain nuclei of live cells and 4µM ethidium bromide to

label nuclei of dead cells. The number of live and dead cells per microscopic field was

counted under a Nikon Eclipse fluorescent microscope using 480 and 520nm filters,

respectively. The level of cell death in the cultures was determined biochemically using

a lactate dehydrogenase (LDH) cytotoxicity assay (Roche, UK). Cell culture

supernatants were analysed after a 2-days’ culture for the presence of LDH. Absorbance

was determined at 490/630nm using the Biotek plate reader.

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2.3 BrdU cell proliferation assay

Cell proliferation was assessed using a 5-bromo-2-deoxy-uridine (BrdU) labeling and

detection kit (Roche Applied Sciences). In brief, cells were labeled with 10 μM BrdU

for the final hour of the 48 hours’ culture followed by fixation and immunostaining for

BrdU incorporation using a specific anti-BrdU antibody. Cells were counterstained with

hematoxylin and the total number of labeled and non-labeled nuclei were counted in 50

independent microscope fields.

2.4 Biochemical alkaline phosphate (AlP) assay

Cells were lysed in 0.1% Triton X-100 and incubated for 10 min in 1 M diethanolamine

buffer (pH 9.8) containing 1mg/ml p-nitrophenyl phosphate (pNPP) at 37° C.

Production of PNP (p-nitrophenol) was quantified spectrophotometrically at an

absorbance of 405 nm using an automatic plate reader.

2.5 Semi-quantitative RT-PCR (sqRT-PCR) analysis

Cells were lysed in RLT buffer containing β-mercaptoethanol followed by RNA

isolation using the RNeasy minikit (Qiagen, UK). Subsequently, 1μg of DNase-digested

total RNA was used for oligo(dT) (Ambion, UK) reverse transcription to generate

single-stranded cDNA using the Omniscript kit (Qiagen, UK). Centrifugal filters

(Microcon) were used to purify and concentrate resultant cDNA. Both RNA and cDNA

concentrations were determined from absorbance values at a wavelength of 260 nm

using a BioPhotometer (Eppendorf, UK). sqRT-PCR assays were performed using the

RedTaq PCR system (Sigma, UK) and the Mastercycler gradient thermal cycler

225

(Eppendorf, UK). Primers were designed from NCBI mRNA sequences using Primer-3

design software (Table 1).

2.6 Immunocytochemistry

MC3T3-E1 were seeded onto multispot microscope slides and incubated for 24h at 37

0C in a humidified 5% CO2 incubator (15,000 cells/well). The adherent cells were fixed

with ice-cold acetone for 5 min followed by rinsing in phosphate-buffered saline (PBS)

containing 1% BSA. Following incubation in 3% H2O2 for 30 min (to block endogenous

peroxidase), the slides were washed in PBS and incubated in 20% normal goat serum

followed by incubation with 2 µg/ml primary polygonal rabbit antibody against GFRα1

(sc10716, SantaCruz) or against RET (sc167; SantaCruz) overnight at 4o

C. Both

antibodies have been shown to specifically recognise the respective protein receptors as

determined by immunoblotting, and have been validated for use in

immunocytochemical staining of cell membrane receptors (see manufacturer’s

datasheets). To demonstrate that the immunocytochemical staining was specific for the

primary antibody, the primary antibody was substituted with 20% normal rabbit serum.

The slides were rinsed in PBS/1% BSA and labeled and stained with biotin-

streptavidin-HRP using a Biogenex detection kit (LP000-UL). The slides were

counterstained with haematoxylin before examination under a Zeiss microscope.

2.7 Data and statistical analysis

226

Data obtained from the WST-1, LDH and ALP assays were corrected for background

values and expressed as percentage of controls. Data were analysed using ANOVA with

Tukey’s posthoc test.

3. Results

3.1 Expression of GDNF, GFRα1 and RET in calvarial osteoblasts

sqRT-PCR analysis revealed that GDNF and its receptors GFRα1 and RET were

expressed in the osteoblast cell line MC3T3-E1 (Fig. 1A). No obvious changes in gene

expression were evident in serum-free cultures as compared to cultures maintained in

serum supplemented media (Fig. 1A). This observation was supported by gel image

analysis (unpublished observations). Immunocytochemical staining of MC3T3-E1 using

specific antibodies against GFRα1 and RET confirmed the presence of these GDNF

receptors (Fig. 1B).

3.2 GDNF stimulates MC3T3-E1 cell proliferation

Addition of GDNF to the osteoblast-like cells did not elicit major changes in the

number of viable cells over a 2-day culture period in medium supplemented with FBS;

however, at 100 ng/ml GDNF a significant, albeit modest decrease in cell number was

evident (82.9% of controls). Conversely, GDNF dose-dependently increased viable

MC3T3-E1 numbers in serum-free cultures (Fig. 2A).

227

Biochemical analysis of alkaline phosphate (ALP), a non-specific marker for early

osteoblast differentiation (19-20) demonstrated that GDNF had no significant effect on

overall ALP activity (data not shown), but following correction for cell numbers a dose-

dependent decrease in ALP levels at increasing GDNF concentrations in serum-free

cultures was evident (Fig. 2B). These data indicated that the reduced relative ALP

activity in GDNF-treated serum-free cultures corresponded with increased cell growth.

To corroborate the WST-1 data, cell proliferation was further analyzed using the BrdU-

incorporation assay. The BrdU data demonstrated that the mitotic activity in serum-free

cultures was greatly reduced compared to the serum-supplemented cultures (Fig. 3).

GDNF had a modest, albeit non-significant, effect on BrdU labeling in serum-

containing cultures (23% reduction compared to controls). However, GDNF

significantly increased the number of BrdU-labeled cells in serum-free cultures by

103.7% (i.e. two-fold increase) demonstrating that GDNF stimulated cell replication

under these conditions (Fig. 3).

3.3 GDNF does not affect osteoblast cell survival

To further investigate whether GDNF influenced cell survival, the live-dead assay was

applied. Results demonstrated that significant cell death occurred in control, serum-free

cultures as compared to serum-supplemented cultures (Fig. 4A). GDNF did not

significantly affect the number of dead cells in either serum-containing or serum-free

cultures suggesting that GDNF did not affect cell death or survival under either

condition (Fig. 4A).

228

These data were corroborated by the biochemical cytotoxicity LDH assay demonstrating

that GDNF did not influence the level of cell death under serum-free conditions (Fig.

4B).

3.4 Receptor-mediated effects of GDNF

To determine whether GDNF affected the cells through its canonical receptors GFRα1

and RET, cultures were treated with specific compounds known to block GDNF

signalling. PI-PLC which hydrolyses the GFRα1 subunits from their

glycosylphosphatidylinositol (GPI)-anchored membrane proteins thereby negating

GDNF signalling via this receptor, abrogated GDNF effects on viable cell numbers in

serum-free cultures (Fig. 5). These data underline an essential role for GFRα1 in the

GDNF effects on osteoblast viability. RPI-1, a competitive ATP-dependent RET kinase

inhibitor, dose-dependently blocked GDNF action indicating that activation of the RET

co-receptor was necessary to elicit GDNF signalling in these cells (Fig. 5).

3.4 Interaction of GDNF with TNFα

Finally this study investigated the effects of GDNF in the presence of the pro-

inflammatory cytokine TNF-α, which is known to have profound effects on bone cells

including MC3T3-E1 osteoblastic cells (21-23). TNF-α dose-dependently increased

viable cell numbers in both serum-containing and serum-free MC3T3-E1 cultures (Fig.

6). Addition of GDNF to the cultures supplemented with TNF-α further promoted

osteoblastic cell growth in these cultures: The stimulating effects of GDNF appeared

229

relatively modest and non-significant in the 10% FBS cultures (Fig. 6A); however,

GDNF synergistically increased cell numbers in the presence of TNF-α in serum-free

cultures (Fig. 6B).

4 Discussion

The current study provides evidence of a direct effect of GDNF on calvarial-derived

osteoblasts suggesting a potential role for this neurotrophic factor in the regulation of

craniofacial bone metabolism. GDNF as well as both of its canonical receptors GFRα1

and RET were shown to be expressed in MC3T3-E1 osteoblastic cells and signaling

through both receptors was needed for GDNF effects on osteoblast cell growth. This is a

novel and interesting finding, as previous studies reported that only GDNF and

GFRα1were present in two human osteosarcoma cell lines (Saos-2, MG63) and primary

bone marrow stromal cells, but not RET (12). Thus this latter work had led to the

conclusion that GDNF signaling in the bone marrow environment involved interaction

with RET-expressing hematopoietic cells (13-14). Previous studies have suggested that

isolated cells from calvarial bones may behave differently than osteoblasts derived from

long bones ((21-23)), which may reflect the mechanistically different processes by

which the different structures in the skeleton develop (i.e. flat bones via

intramembranous bone formation, whereas long bones through the process of

endochondral bone formation; (27)). Moreover, it is worth noting that a significant part

of the craniofacial skeleton originates from neuronal crest ―ecto-mesenchymal‖

progenitor cells and therefore the calvarial osteoblasts may therefore exhibit a different

molecular repertoire and cell behavior than mesodermal/mesenchymal-derived bone-

forming cells present in long bones (24-26). RET is considered important for

230

development and differentiation of neural crest-derived tissues, including cranial tissues

(28-30). GDNF was shown to be co-expressed with GFRα1/2 and RET in dental

epithelial and mesenchymal cells during tooth development (31,32). Our recent studies

indicated that mesenchymal/stromal cell cultures derived from dental pulp also

displayed co-expression of GDNF and GFRα1/RET (33). RT-PCR analysis suggested

that the calvarial osteoblasts expressed GDNF and the receptors GFRα1 and RET in

serum-free cultures to a similar degree as cells maintained in serum-supplemented

media.Previous studies demonstrated gene expression of both GDNF receptor

components GFRα1 and RET in adrenal medullary cells and glial cells cultured in

serum-free medium (34, 35).Interestingly, addition of serum upregulated the

transcription levels of GFRα1 and RET in these cells (34, 35). Further quantitative RT-

PCR is recommended to evaluate and substantiate the effects of culture conditions on

gene expression in osteoblasts; notwithstanding our observations indicate that the

calvarial osteoblastic cell line expressing the primary GDNF receptors represents a

suitable model to investigate GDNF signaling in either serum-containing or serum-free

culture media.

This study demonstrates that GDNF stimulated MC3T3-E1 cell proliferation in

serum-free conditions, which corresponded with a concomitant inhibitory effect on

relative ALP activity, a marker of early osteogenic differentiation. Moreover, our data

indicate that GDNF at high concentrations exerted a cell growth curbing effect on

osteoblasts maintained in serum-supplemented culture medium; an effect that appeared

to be associated with a slight increase in ALP activity (Fig. 2). It is well established that

cell differentiation is often inversely related with mitotic activity and our observations

indicate that GDNF effect on calvarial osteoblasts mainly involves an action on cellular

231

proliferation with a concomitant inverse effect on immediate differentiation.

Interestingly, GDNF/RET signaling has been shown to be responsible for the anti-

mitotic action in an embryonic neural precursor carcinoma cell line (36). This effect

mediated by p27kip1

was suggested to be a mechanism by which GDNF regulates cell

growth to initiate terminal differentiation (36). Long-term culture experiments will be

required to investigate in further detail the role of GDNF in osteogenic proliferation,

differentiation and bone formation.

The mitotic and cell survival actions of GDNF through GFRα1/RET signaling

are well documented in the literature underscoring the multifunctional role of GDNF in

tissue maintenance, repair and regeneration (1-3). Shi et al (37) described that GDNF

promoted mesenchymal stem cell migration and survival; a mechanism by which GDNF

may deliver renoprotection and kidney repair. The physiological implications of our

findings that GDNF stimulated calvarial osteoblast proliferation under serum-deprived

conditions are as yet unclear, but may allude to a novel role of this neurotrophic factor

in bone remodeling and repair. GDNF is considered a member of the TGFβ superfamily

as it has a partial amino-acid sequence homology and similar structural confirmation to

TGF (38), which comprise growth factors including bone morphogenetic proteins

pivotal in regulation of bone development, metabolism and repair. TGF1 can have

diverse and multiple effects in different cell systems; notably this signaling molecule

has been ascribed a central role in bone cell recruitment, proliferation and

differentiation (39). Interestingly, TGF1 induces translocation of GFR1 to the plasma

membrane thereby enabling GDNF signaling through this receptor (40). It is also worth

noting that GDNF is a potent inducer of the nuclear transcription factor, murine GDNF

232

inducible factor (mGIF) which is homologous to the human TGFβ inducible early gene

(TIEG) (41). TIEG expression which has been suggested to play a pivotal role in the

regulation of osteoblast differentiation (42) is highly induced in human osteosarcoma

cells as well as immortalized human fetal osteoblasts following treatment with TGFβ

(43). It would be fascinating to explore whether GDNF signaling in calvarial

osteoblasts is related to induction of TIEG/mGIF.

Considering that GDNF is produced by bone marrow stromal cells as well as

osteoblasts (9, 44, 45), it is tempting to speculate that GDNF in conjunction with other

auto- and paracrine factors may be involved in the regulation of osteoblast recruitment

in bone growth and remodelling. Indeed, this study demonstrated that GDNF cooperated

with the cytokine TNF-α to stimulate osteoblastic cell growth suggesting an interaction

between GDNF and TNF-α signalling pathways in osteoblasts. TNF-α has been ascribed

a multifunctional role in bone metabolism (22); TNF-α effects may involve a pro-

resorptive (osteoclastic bone degradation) action during inflammatory conditions (21),

but TNF-α has also been recognised as an anabolic cytokine stimulating osteogenic

migration, proliferation and differentiation (46-50). Interestingly, TNF-α has been

shown to have neuroprotective capabilities which in part may be dependent on

induction of cytoprotective neutrotrophic growth factors such as GDNF (51-53).

Moreover TNF-α was reported to induce GDNF in chondrocytes underscoring a

potential role for GDNF in skeletal cells under pro-inflammatory conditions (54).

Further research is warranted to explore the precise role and mechanistic interaction of

these pleiotrophic signaling molecules in bone remodeling and their potential

therapeutic use in bone regeneration and repair.

233

In conclusion, this is the first study to report that the neurotrophic factor GDNF

is able to influence the proliferation of calvarial osteoblasts via its canonical receptors

GFRα/RET expressed on these cells highlighting a novel regulatory pathway in

craniofacial bone physiology.

Acknowledgements

This study was supported by a University of Birmingham School of Dentistry PhD

research grant. We are grateful to Amgen Ltd (Thousands Oaks, USA) for the supply of

the recombinant GDNF.

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Tables and figures

Table 1. PCR primer sequences and annealing temperature (Tm)

Gene

symbol

Primer sequence (5’ to 3’) Genbank

Accession no.

Product

size

Tm

GAPDH F-CCCATCACCATCTTCCAGGAGC

R-CCAGTGAGCTTCCCGTTCAGC NM017008 450bp 60

GDNF F-AGAGGAATCGGCAGGCTGCAGCTG

R-AGATACATCCACATCGTTTAGCGG NM019139 337bp 60

RET

F-TCAGGCATTTTGCAGCTATG R-TGCAAAGGATGTGAAAGCAG

NM001110099 393bp 62.5

GFRα1 F-AATGCAATTCAAGCCTTTGG R-TGTGTGCTACCCGACACATT

U59486 218bp 60

240

Fig. 1. (A) Representative RT-PCR gel images demonstrating the presence of

transcripts for GDNF and its receptors GFRa1 and RET in MC3T3-E1 calvarial

osteoblasts cultured

in serum-supplemented (10% FBS) or serum-free culture medium for 2 days. C6 glioma

cells were used as positive control, and glyceraldehyde-3-phosphate dehydrogenase

(GAPDH) as control, housekeeping gene. Respective cycle number used for GDNF,

GFRa1, RET and GAPDH were 35, 45, 45 and 25. (B) Immunocytochemical staining of

MC3T3-E1 for GFRa1 and RET. Positive staining (brown) for both receptors was

clearly evident in the MC3T3-E1 cells, whilst immunostaining was absent in controls

(specific

primary antibody was substituted with non-immune rabbit serum). (For interpretation of

the references to colour in this figure legend, the reader is referred to the web

version of this article.)

241

Fig. 2. (A) Effect of GDNF on viable MC3T3-E1 cell number in 2-day serum

supplemented

(10% FBS) or serum-free (0.1% BSA) cultures as assessed by WST-1.

Results are expressed as percentage of controls (mean ± SD; n = 4). (B) Effect of

GDNF on alkaline phosphatase (AlP) activity in osteoblast cultures after 2 days of

culture. Results represent relative AlP activity corrected for cell number (percentage

of controls; mean ± SD; n = 4). ⁄P < 0.05, ⁄⁄P < 0.01 versus control values.

242

Fig. 3. BrdU incorporation in 2-day MC3T3-E1 cultures in serum-supplemented

(10% FBS) or serum-free (0.1% BSA) cultures. The results show the proportion of

cells labeled positively for BrdU (Li: labeling index). Significantly different from

controls: ⁄P < 0.05.

*

243

Fig. 4. Effects of GDNF on cell death in MC3T3-E1 cultures. (A) The number of dead

cells as determined by the live/dead assay in cultures treated with 100 ng/ml GDNF.

Results show percentage of dead cells in 1- and 2-day cultures (mean ± SD; n = 3).

(B) Relative LDH levels after 2-days culture in serum-containing or serum-free

medium cultures supplemented with 100 ng/ml GDNF (mean ± SD; n = 3). Results

are mean ± SD (n = 3–4).

244

Fig. 5. Effects of GDNF receptor inhibitors on GDNF-stimulated MC3T3-E1 cultures.

(A) Effect of phosphatidylinositol-specific phospholipase C (PLC) on viable cell

number in cultures supplemented with 100 ng/ml GDNF. (B) Effects of the RET

kinase inhibitor, RPI-1, on viable cell number in 2-day GDNF-treated cell cultures.

Results are percentage of control values as determined by the WST1 assay

(mean} SD of 6.8 replicates). Data from GDNF control cultures (without inhibitor)

were significantly different from controls without GDNF (P < 0.01). ..P < 0.01 versus

GDNF cultures.

** **

245

Fig. 6. Osteoblastic MC3T3-E1 cell numbers after 2-days’ culture in the presence of

increasing concentrations of TNF-a with or without 100 ng/ml GDNF in media

supplemented with 10% serum (A) or in serum-free cultures (B).Viable cell numbers

were assessed using the WST-1 assay; results are expressed as percentage of

controls (mean ± SEM; n = 3). ⁄P < 0.05, ⁄⁄⁄P < 0.001 GDNF-supplemented cultures

versus corresponding TNF-a controls.

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